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Cytoplasmic dynein motors are localized to the cytoplasmic face of the er component of es in rat sertoli… Kimel, Gil Howard 2000

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CYTOPLASMIC DYNEIN M O T O R S ARE LOCALIZED TO T H E CYTOPLASMIC F A C E O F T H E E R C O M P O N E N T OF E S IN R A T SERTOLI C E L L S by Gil Howard Kimel B.Sc, McGill University, 1996 A THESIS SUBMITTED IN PARTIAL FULFILLMENT O F T H E REQUIREMENTS FOR T H E DEGREE O F MASTER O F SCIENCE in The Faculty of Graduate Studies (Department of Anatomy) We accept this thesis as conforming to the required standard The University of British Columbia March 2000 © Gil Howard Kimel, 2000 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. The University of British Columbia Vancouver, Canada Department DE-6 (2/88) Abstract In vitro motility assays indicate that minus-end directed microtubule dependant motors are attached to specialized junction plaques (ectoplasmic specializations) that occur in regions of Sertoli cells attached to spermatids. These plaques are characterized by a layer of hexagonally arrayed actin filaments situated between and linked to a cistern of ER and the plasma membrane. In this study, I use immuno-EM and actin-severing (gelsolin) experiments to test the hypothesis that cytoplasmic dynein is localized to the ER. For immuno-EM, perfusion fixed rat testis were probed for the intermediate chain of cytoplasmic dynein (IC74) before (mechanically fragmented tissue) or after (intact tissue) embedding and compared with controls. Labeling was clearly evident along, although not restricted to, the cytoplasmic face of the ER. For the gelsolin experiments testicular fractions enriched for spermatid/junction complexes were incubated with and without the gelsolin enzyme, which was used to artificially disassemble the junction plaque and release the ER, centrifuged, and the supernatants compared by western blot analysis for GRP94 (a marker for ER), IC74 (Pfister and Babco), and DHC1a (Asai). All three probes reacted more strongly with appropriate bands from gelsolin treated supernatants than with corresponding bands from controls. The data are consistent with the conclusion that a cytoplasmic dynein is associated with ectoplasmic specializations and that the motor is anchored to the cytoplasmic face of the ER component of the plaque. Because Sertoli cell microtubules have their minus-end located apically, a cytoplasmic dynein may be the motor associated with moving spermatids to the apex of the epithelium prior to sperm release. TABLE OF CONTENTS Dedication Abstract ii Table of contents I i i List of figures vi List of abbreviations v i i i Acknowledgments IX CHAPTER 1. General Introduction 1 The Testis: General Organization 4 Spermatogenesis & The Sertoli cell 8 The Proliferative Phase 8 The Meiotic Phase 9 The Spermiogenic Phase 9 The Sertoli cell 10 The Ectoplasmic Specialization 12 Historical Perspective 12 Other Molecules Associated with ESs 13 Structure of Ectoplasmic Specializations 18 The Five Domains of the Ectoplasmic Specialization 21 Cytoplasmic Domain 21 Endoplasmic Reticulum Domain 21 The Ectoplasmic Domain 22 The Plasma Membrane-related zone. 22 The Actin Zone 23 The Endoplasmic Reticulum-rel. zone 23 Plasma Membrane Domain 23 Extracellular Domain 24 Microtubules in General 25 Tubulin 25 Microtubule Structure 25 MAPS 26 Microtubules in Sertoli Cells 26 Polarity of Microtubules in Sertoli Cells 26 The Interaction of Microtubules and Motor... 27 Proteins Motor Proteins in General 29 Dynein 30 Brief History of Dynein Motors 30 i i i Cytoplasmic Dynein 31 Intro, to the Role of Cytoplasmic Dynein.. 31 Structure of Dynein 33 Dynein Heavy Chain 33 Other Elements of Dynein Motor Protein.. 34 The Dynactin Complex 34 Microtubule-Binding Stalks 35 Control of Dynein Mechanics 37 Cytoplasmic Linker Proteins (CLIPs) 37 CLIP-170 38 Microtubules Associate with ER 40 Cytoplasmic Dynein and Apically- Directed 41 Spermatid Translocation Brief Introduction to the Motility Hypothesis 45 The Thesis Problem 48 Hypothesis 48 Approach 48 Rationale 48 CHAPTER 2. Cytoplasmic Dynein is Localised to the Cytoplasmic Face of ER Component of ES: Morphological Evidence 51 Introduction 51 Materials and Methods 52 Animals 52 Reagents and Antibodies 52 Orchotomy 53 Electron Microscopy-Epon 54 Repetition of Experiments 56 Electron Microscopy-Lowicryl/Unicryl 56 Epon vs. Lowicryl 58 Results 61 Electron Microscopy 61 Epon Immunogold Study 61 Lowicryl Immunogold Study 62 Discussion 76 IC74 Immuno-EM 81 Control of Spermatid Translocation 82 iv CLIPS may link motors to ES 87 Quantity & Density of Gold Particles 95 CLIPs and Spermatid Translocation 95 Limitations of EM Study. 96 CHAPTER 3. Cytoplasmic Dynein is Localised to the ER Component of ES Biochemical Evidence 97 Introduction 98 Materials and Methods 100 Testis Prep, for Gelsolin Expt 100 Gelsolin 101 ER Enrichment Protocol. 102 Gelsolin Experiment 102 Western Analysis 103 Repitition of Western Analysis Expts 106 Results 108 GRP94 108 IC74 108 DHC1a 108 Discussion 114 IC74-lmmunoblots 114 Limitations of Biochemical Study 115 DHC1 a- Immunoblots 117 GRP94 118 Overall Conclusions 119 Bibliography 1 2 0 V List of Figures: Page FIGURE 1 Schematic diagram of a Sertoli cell with associated germ 7 cells illustrating the compartmentalization of the germinal epithelium by tight junctions between adjacent Sertoli cells (Fawcett, 1970). FIGURE 2 Electron micrograph of a typical spermatid displaying an ES 15 FIGURE 3 Diagram illustrating the arrangement of actin filaments, 17 intermediate filaments and microtubules in Sertoli cells (modified from Vogl et al., 1991). FIGURE 4 Diagram of the structure of the ectoplasmic specialization at apical 20 and basal sites in the Sertoli cell (Vogl et al., 1991). FIGURE 5 Electron micrographs ((A)cross section, (C) longitudinal) of 44 spermatids showing the relationship with the ES and (B) schematic diagram of the translocation hypothesis. FIGURE 6 The spermatid translocation model 47 FIGURE 7 Electron micrographs of IC74 immunogold-EM study (Epon). 65 FIGURE 8 Electron micrographs of IC74 immunogold-EM study (Epon). 67 FIGURE 9 Electron micrographs of IC74 immunogold-EM study (Epon). 69 FIGURE 10 Electron micrographs of IC74 immunogold-EM study (Lowicryl). 71 vi FIGURE 11 Electron micrographs of IC74 immunogold- EM study (Unicryl). 73 FIGURE 12 Electron micrographs of IC74 immunogold- EM study (Unicry). 75 FIGURE 13 Schematic diagram of the proposed relationship between the ES 80 and Cytoplasmic dynein. FIGURE 14 Schematic four panel diagram illustrating a possible mechanism 90 of directional control of spermatid translocation. FIGURE 15 Schematic four panel diagram illustrating a possible mechanism 92 of directional control of spermatid translocation. FIGURE16 Schematic four panel diagram illustrating a possible mechanism 94 of directional control of spermatid translocation. FIGURE 17 Schematic diagram summarizing the gelsolin experiment. 111 FIGURE 18 Western blot analysis of gelsolin and non-gelsolin treated tissue 113 probed with an anti-GRP, and an anti-IC74 antibody, and an anti-DHC1a antibody. Fluorescence images are also present to demonstrate gelsolin mediated digestion of actin layer of ES. v i i List of Abbreviations: ATP Adenisine 5 '-Triphosphate BSA Bovine Serum Albumin D i O C 6 3,3 '-dihexylocarbocyanine Iodide DTT Dithiothreitol EGTA Ethylene Glycol bis[succininidyl succinate] EM Electron Microscopy ER Endoplasmic Reticulum ES Ectoplasmic Specialization GTP Guanosine 5 '-Triphosphate HRP Horse Radish Peroxidase IC74 Cytoplasmic Dynein Intermediate Chain (74 kDa) KCI Potassium Chloride NaCI Sodium Chloride N a 2 H P 0 4 Disodium Phosphate NGS Normal Goat Serum M g C I 2 Magnesium Chloride PBS Phosphate Buffered Saline PIPES Piperazine-N,N'-bis[2-ethanesulfonic acid] PMSF Phenylmethylsulfonylf louride PVDF Polyvinylidene Fluoride SDS-PAGE Sodium Dodecyl Sulfate-Poly Acrylamide Gel Electrophoresis TPBS Tween-Phospahte Buffered Saline Tris-Hcl (Tris[hydroxymethyl]aminomethane)-Hydro Chloride Tween-20 Polyoxylethylenesorbitan Monolautate Viii Acknowledgments I would like to thank my supervisor and advisor, Dr. A. Wayne Vogl, for his encouragement, support and enthusiasm throughout my M.Sc. project. I also thank him for giving me the freedom to learn and to experiment with the scientific techniques that I was interested in and also to pursue several upper-level anatomy courses. I also learned from and really enjoyed being a teaching assistant in the medical gross anatomy course that he directed. I will remember with much fondness some of the lively discussions and debates that we shared! I would like to thank Dr. Tim O'Connor and Dr. Linda Matsuuchi, both members of my supervisory committee, for many very helpful discussions especially with regards to the biochemical studies in this project. I would also like to thank them for the improvements that they made to this manuscript. In addition, I would like to thank the Roskelley Lab for helping me with some of the technical aspects of the Westerns blots. I would also like to thank Candace Hoffmann for helping me with the statistical analysis. Thanks also to Drs. Asai and Pfister for providing me with the anti-dynein antibodies. Cheers to Jonathon Wilson, Roseanne Mclndoe, Jullin Green, and Elaine Humphrey. I would like to extend a special thanks to Dr. Charles Slonecker for supervising me for my directed studies course and for mentoring me throughout my studies. Thanks also to Drs. Emerman, Weinberg and Hollenberg for advising throughout my studies. I would like to thank my Dad, Judith and Debbie for their love, support and guidance and also for supporting my decision to move to Vancouver in the first place! Lastly, I would like to thank my very close and special friends with whom I have shared so much with over the last few years. ix Funding for this study was provided by the Medical Research Council of Canada (Grant # MA-13389) awarded to Dr. A.W. Vogl. dedicated to Ruth Kimel Xi' CHAPTER 1 GENERAL INTRODUCTION 1 In this thesis I present evidence that is consistent with the hypothesis that cytoplasmic dynein is anchored to the cytoplasmic face of the endoplasmic reticulum (ER) component of ectoplasmic specializations (ES) that occur in Sertoli cell regions adjacent to spermatid heads. During mammalian spermatogenesis, a conspicuous, dynamic and microtubule-dependent (Vogl, 1996) event occurs in the seminiferous epithelium- the translocation of spermatids. Spermatids are translocated in two opposite and parallel directions. Spermatids are initially shuttled basally, towards the Sertoli cell nucleus and then apically, towards the lumen of the seminiferous tubule. The function of this translocation event is not known; however the surface area contact between elongate spermatids and Sertoli cell crypts is greatly increased when spermatids are translocated to the basal region of Sertoli cells. Thus, it has been proposed that optimisation of intercellular communication may be the purpose for the translocation event (Beach and Vogl, 1999). In brief, my hypothesis is that cytoplasmic dynein motors are localized to the cytoplasmic face of the ER component of a specialized junctional plaque (ectoplasmic specialization) found in the testis. Although a recent study (Miller et al., 1999) has demonstrated, with immunofluorescence light microscopy, that cytoplasmic dynein (intermediate chain- IC74) is localized to the area surrounding the spermatid head, there are no studies that have precisely 2 localized, by means of electron microscopy, the dynein motor protein at the level of the ES. In this thesis, I present morphological and biochemical data indicating that cytoplasmic dynein is localized to the cytoplasmic face of the ER component of ES. This study is an important contribution to the field of reproductive biology and cell biology since it proposes the involvement of cytoplasmic dynein, a microtubule dependant motor, in spermatogenesis. A complete understanding of the mechanism responsible for the spermatid translocation event may, in the future, present an opportunity to develop drugs that could influence spermatogenesis. 3 The Testis: General Organization The testis, incapsulated by a clear and tough fibrous sheath, is situated in the scrotum. Once the testis is removed from the scrotum and dissected from its fibrous sheath, the remaining mass of seminiferous tissue may be divided into two main compartments, namely: the interstitial compartment and the seminiferous tubule compartment. Present within the interstitial compartment are the vascular and lymphatic systems and Leydig cells. Unlike blood vessels in most glands, testicular capillaries are non-fenestrated (Russell et al., 1990). Leydig cells, which are responsible for the production of testosterone and other steroids, are very abundant in the interstitial compartment (Russell et al.,1990). Seminiferous tubules house Sertoli cells and associated spermatogenic cells which together constitute the epithelium. Sertoli cells are responsible for 'supporting' the differentiating spermatogenic cells. Seminiferous tubules are highly convoluted tubes and generally run in the direction of the long-axis of the testis. In rats, their walls are composed of various cellular and acellular (extracellular matrix) layers. Directly surrounding the epithelium of the tubules is a basal lamina which is surrounded by a collagen layer, myoid cells another collagen layer and finally lymphatic channels which are covered by a layer of endothelial cells. The myoid cells, which have "muscle-like" properties, are thought to be important for the propulsion of sperm through the seminiferous tubules (Tripiciano et al., 1999). 4 Myoid cells also are thought to structurally support the seminiferous tubules (Russell et al., 1990). 5 Figure 1. - Relative Positioning of Various Stages of Sperm Cells with Respect to a Sertoli Cell. A diagram from Dym and Fawcett (1970), depicting the compartmentalization of the germinal epithelium by tight junctions (asterisk) between adjacent Sertoli cells. The tight junctions between two adjacent Sertoli cells separate the basal compartment, containing the spermatogonia and early preleptotene spermatocytes, from the adluminal compartment, containing the spermatocytes and spermatids. 6 7-Spermatogenesis Spermatogenesis is the process whereby spermatogonia differentiate into spermatozoa. Spermatogenesis in rat lasts 57 days (Leblond and Clermont 1952). This process may be divided into three distinct phases. 1 1. The proliferative phase is characterised by rapid and successive divisions of the spermatogonia. 2. The meiotic phase is characterized by the formation of haploid spermatids. 3. The spermiogenic phase is characterized by the maturation of spermatids into spermatozoa which are structurally prepared for fertilization (Russell et al.1990). The Proliferative Phase During the proliferative phase a renewal of the spermatogonia cell population occurs. Immature spermatogonia undergo numerous mitoses to build a large population of cells that will subsequently undergo meiosis and differentiate into mature sperm cells. The most immature spermatogonia are resistant to common insults (such as temperature fluctuations) and will often survive when other stages of germ cells die. This resistance is crucial since complete destruction of stem cells would lead to sterility. Spermatogonia in the proliferative phase may be grouped into one of the three following divisions depending on the amount of chromatin lying along the inner aspect of the nuclear envelope: (1) Type A, (2) Intermediate, and (3) Type B (Russell et al.1990). 8 The Meiotic Phase The meiotic phase is characterized by cell division of spermatogonia to form primary spermatocytes. More specifically, type B spermatogonia divide to form preleptotene cells, which are the last cells to pass through the S-phase of the cell cycle. During meiosis, chromosomes are recombined and the genetic material is halved. Once the meiotic divisions are completed the germ cell number is quadrupled. Prophase of the first meiotic division lasts approximately three weeks. During prophase the cells and their nuclei increase in size, however, this increase in size becomes much more pronounced in late prophase. The meiotic prophase is subdivided into the following sub-stages: (1) preleptotene, (2) leptotene, (3) zygotene, (4) pachytene, (5) diplotene. These subdivisions are based on morphological changes of the nucleus (Russell et al.1990). The Spermiogenic Phase During the spermiogenic phase the maturation of spermatids to spermatozoa occurs. The first morphological sign of the spermiogenic phase is the appearance of a flagellum. Once the centriole pair migrates to the cell surface, one of the two centrioles forms an axoneme. This structure contains microtubules which cause the plasma membrane to protrude from the cell. Later on in the maturation process, the flagellum, undergoes the addition, to itself, of a middle, principal and end piece. Mitochondria are recruited from the cytoplasm to form a helical pattern around the middle piece. Interestingly, even though spermatids acquire a flagellum while they are in the testis; spermatids 9 are immotile until they are expelled from the testis. In summary, during this phase, spermatids acquire an acrosome, the nucleus condenses, and the cell acquires its species specific shape (Russell et al. 1990). The Sertoli Cell The Sertoli cell, named after its discoverer, Enrico Sertoli, is a very complex cell that interacts with and supports maturing germ cells. During puberal development, Sertoli cells cease to divide and it is thought that their population remains constant throughout the life span of the animal. The Sertoli cell is a polarized columnar epithelial cell that extends, approximately 100 um (in rat), from the base (basal lamina) towards a central point in lumen of the seminiferous tubule. The nucleus of the Sertoli cell is indented and characterized by a tripartite nucleolus. Serial section reconstruction of the Sertoli cell revealed its three-dimensional structure and thus, demonstrated that it does not form a syncytium with other Sertoli cells. The approximate volume of a Sertoli cell is 6000 um3, whereas its surface area is about 12,000 um 2 (Russell et al. 1990). Sertoli cells are attached to the basal lamina via hemidesmosomes; to adjacent Sertoli cells by desmosomes, gap junctions, and tight junctions (blood testis barrier) and to germ cells via ESs, gap junctions and tubulobulbar complexes. Through these junctions Sertoli cells maintain the structural integrity of the epithelium and also provide for cell to cell communication. Gap junctions in particular are thought to provide a means for Sertoli cell-germ cell communication. Tubulobulbar complexes anchor the 10 germ cells to Sertoli cells and eliminate the germ cell cytoplasm before sperm release (Russell et al. 1990). Sertoli cells are divided into two permanent compartments (basal and adluminal) and one transient (intermediate- between basal and adluminal) compartment. The basal compartment contains immature germ cells (spermatogonia and early spermatocytes). The adluminal compartment contains the more mature spermatids and spermatozoa. Since cells in the basal compartment are not protected by the blood-testis barrier they have free access to substances diffusing from the lymphatic and vascular systems. Tight junctions (Figure 1, asterisk), forming the intermediate compartment (blood-testis barrier), disassemble to allow for leptotene cells to transit through this zone and then reform once the cell enters the adluminal compartment (Russell et al. 1990). Sertoli cells secrete fluid apically to maintain the seminiferous tubule lumen. This luminal fluid transports the non-motile (spermatids only become motile in the epididymis) sperm within the duct system of the male reproductive tract. Sertoli cells are also actively involved with spermiation- the process whereby sperm are released to the lumen of the tubule. Sertoli cells may also be phagocytic of germ cells that do not develop properly and the tubulobulbar complexes. Sertoli cells nourish germ cells by delivering substances, such as lactate (which is produced by the Sertoli cell from substrates originating in the vascular system) directly to the germ cells (Russell et al. 1990). 11 Ectoplasmic Specializations ESs are specialised structures that occur in areas of contact between neighbouring Sertoli cells and between Sertoli cells and developing spermatids. Thus, these specialized junctions are located both in apical and basal regions of the Sertoli cell. At apical sites, ESs lie directly opposed to the plasma membranes of the acrosomal area of spermatids. At basal sites, ESs form contact points (tight junctions) between two adjacent Sertoli cells. Historical Perspective The first morphological description of ESs, based on studies of the junctional complex linking adjacent Sertoli cells, were provided by Brokelmann (1961, 1963) and Nicander (1963). After further study this structure was given the name 'junctional specialization' (Flickenger & Fawcett, 1967). Subsequent to this, a similar structure was then localised to the Sertoli-spermatid junction and termed 'ectoplasmic specialization' (Russell, 1977). Toyama, who, in 1976 showed that meromyosin binds to a component of the ES concluded that there were actin filaments localized to this junction and speculated that the ES was contractile (Toyama, 1976; Vogl et al., 1983). Subsequent to this, Gravis demonstrated that there were ATPases in close association with micro-filaments of the ES and that these ATPases may be associated with motility (Russell, 1977). However, now it is generally accepted that actin bundles in ESs do not have contractile potential and are most likely associated with a skeletal or structural function (Vogl and Soucy, 1985). 12 Russell studied the association of ESs with spermatids. He found that ESs were never seen in association with preleptotene, leptotene, zygotene or early pachytene spermatocytes. In fact, according to Russell, the first stage to be consistently associated with ESs, were late pachytene spermatocytes. Russell also postulated that the ES may be more of a supportive pocket for spermatids rather than an adhesion-type junction. In his study he suggested that the ES conforms to the shape of the spermatid in a loose rather than in a binding fashion (Russell, 1977). Russell also compared the appearance of the Sertoli cell- Sertoli cell ESs with the Sertoli cell- spermatid ESs and found that the endoplasmic reticulum of the latter displayed much fewer ribosomes. In fact, very few if any ribosomes were seen on the membranes of the ER at Sertoli cell- spermatid ESs. He also reported that, except for some 'fuzzy material' there did not seem to be any type of 'connecting material' between two adjacent Sertoli cells. Russell reasoned that the 'fuzzy material,' which was visible on electron micrographs, was most probably the cell coat (glycocalyx) (Russell 1977). Other Molecules Associated With ESs In the last twenty years a diverse group of molecules have been localized to ESs of Sertoli cells. For instance a-actinin (Franke et al., 1978), myosin Vila (Hasson et al., 1995), fimbrin (Grove et al., 1989), a6p1 integrin (Pfeiffer et al., 1991) and most recently espin (Bartles et al., 1996). The function of all of these molecules at the level of the ES remains to be clarified. 13 Figure 2. - EM of a Typical Spermatid. An electron micrograph (Epon embedded) of a cross section of an isolated elongate spermatid with its associated ES junctional complex, Magnification: X 36,400, bar = 0.5 um. 14 Figure 3. - Organization of Sertoli Cell Cytoskeleton and its Relationship with a Spermatid. A schematic diagram modified from Vogl et al. (1991) illustrating how actin filaments, intermediate filaments, and microtubules are organised at apical ESs. In particular note the close relationship between the ES junction and the spermatid and between the parallel microtubules and the cytoplasmic face of the ER. The basal ES is shown as a band, at the level of the Sertoli cell nucleus, surrounding the cell (actin of basal ES is labelled). 16 Structure of Ectoplasmic Specialization (ES) The general structure of the ES is as follows. Starting at the most cytoplasmic (labelled as cytoplasmic domain on Figure 4) aspect of the junction plaque, the first component of the ES is the ER. The ER comes into direct contact with two ES associated structures: microtubules (cytoplasmic side of ER) and the actin layer (ectoplasmic side of ER). Between the ER and the actin zone is an area known as the 'ER related zone' and between the actin zone and the plasma membrane of the Sertoli cell is the 'plasma membrane related zone'. The plasma membrane of the spermatid is immediately adjacent to the plasma membrane of the Sertoli cell (Vogl et al., 1991). The ES has been divided into five domains (Figure 4), the Cytoplasmic Domain, the ER Domain, the Ectoplasmic Domain, the Plasma Membrane Domain and the Extracellular Domain (Vogl et al., 1991). 18 Figure 4. - The Domains of the Ectoplasmic Specialization and their Relationship with the Spermatid. A schematic diagram from Vogl et al., (1991) illustrating the structural organization of the ES both at apical regions (ES junction between Sertoli cell and spermatid) and at basal regions (ES junction between two adjacent Sertoli cells) in the mammalian seminiferous epithelium. 19 Domains Cytoplasmic Domain Intermediate! Filaments Microtubule - ER Related Zone Endoplasmic T Reticulum , — P M Related Zone Sertoli Cell Actin Zone! Ectoplasmic Plasma Membrane - Extracellular Spermatid [_ Ectoplasmic Specialization Apical Junction |- Between Sertoli Cell And Spermatid Basal Junction Between Adjacent Sertoli Cells Sertoli Cell Sertoli Cell The Five Domains of the Ectoplasmic Specialization (1) Cytoplasmic Domain The cytoplasmic domain is located just cytoplasmic to the ER and most likely contains structures that link the ER with microtubules (such as microtubule dependent motors) and perhaps even the intermediate filaments. Microtubules located in this area are abundant and it has been suggested that they play a role in the apical translocation of spermatids during spermatogenesis (Redenbach & Vogl, 1991; Vogl, 1996; Beach & Vogl, 1999). This suggestion is supported by the fact that microtubules are oriented parallel to the long axis of the Sertoli cell crypt and also because linkages between the ER and the microtubules have been visualised ultrastructurally (Russell, 1977). Other studies have suggested that these linkages may, in fact, include motor proteins (Vogl, 1996). (2) Endoplasmic Reticulum Domain The endoplasmic reticulum domain is a specialized, ER-like, double membrane structure unique to Sertoli cells. Although this domain is similar, morphologically, to the well-studied endoplasmic reticulum system present in all eucaryotic cells it is not known whether this ER performs functions similar to eucaryotic ER in general, i.e. mediation of selective transfer of molecules between this compartment and the cytosol or the biosynthesis of proteins and lipids. Fenestrated ER cisterns (the ER of ES has 'windows') surround spermatid heads (Figure 3) by lining the crypts in which spermatid heads are 21 located. The ER has not been extensively studied both in terms of precise morphology or specific function, however, it has been suggested that this domain may play a role in the local regulation of calcium levels. Franchi et al., (1985), have treated the ES with trifluoroperazine, a calmodulin inhibitor, and have reported that this compound severely disrupts actin networks and junctions in rat testis. The possibility that local calcium concentrations may be controlled by the ER is interesting and worth further study. (3) The Ectoplasmic Domain This domain, composed mostly of actin filaments and related actin binding proteins, is situated between the ER and the Sertoli cell plasma membrane. This domain is further sub-divided into three zones, namely: (a) Plasma Membrane- related zone (b) Actin Zone (c) Endoplasmic Reticulum-related zone. (a) The Plasma Membrane- related zone Linkages are present between the plasma membrane of the Sertoli cell and the actin zone. These linkages have been identified morphologically and appear as fibrillar strands or as clumps of amorphous material (Grove et al., 1986). Also, based on immunological studies it appears that vinculin is in this zone. Vinculin is a 130 kD protein that is often associated with actin adhesion sites in general. Although unrelated to ESs, ZO-1 (Zonula occuldens-1) is a peripheral membrane protein that may be associated specifically with tight junctions found in basal regions of this zone (Byers et al., 1988). 22 (b) The Actin Zone The actin zone, composed of a layer of actin filaments is usually four to six filaments thick. The arrangement of these filaments is in highly organised bundles that are hexagonally packed and have a uniform polarity. According to morphological studies the space between adjacent filaments is 10-11 nm. Linkages between adjacent filaments can often be detected with electron microscopy. In apical areas of Sertoli cells, actin filament bundles change dramatically throughout spermatogenesis in order to accommodate the shape change of associated spermatids. In rat, at apical ES sites, actin filaments generally run parallel to the long axis of the spermatid head. Actin filaments also outline the Sertoli cell at its basal face. (c) The Endoplasmic Reticulum- related zone This zone is located between the actin zone and the ectoplasmic face of the ER. When spermatids are mechanically torn from Sertoli cells, the ES (including the ER component) usually remains firmly anchored to the spermatid. The strength of this membrane system suggests that there are linkages between the ectoplasmic face of the ER and the actin zone to resist these tearing forces. (4) Plasma Membrane Domain The Sertoli cell plasma membrane component of the ES has not been extensively studied; however, it is suspected that this membrane may have specialised molecules that are essential for intercellular (Sertoli cell -Spermatid) adhesion. It is thought that actin filaments may be linked to cell 23 adhesion molecules (CAMs) and thus, may be involved with intercellular adhesion (Vogl et al. 1991). pi integrin (Pfeiffer et al., 1991) and a6 integrin (Salanova et al., 1995) have been identified at both apical and basal sites of Sertoli cells as transmembrane adhesion molecules. (5) Extracellular Domain The extracellular domain is defined as the space between the plasma membrane of the Sertoli cell and the plasma membrane of the spermatid. Linkages between these two adjacent membranes have been observed with electron microscopy, however, only at apical sites (Russell et al., 1985). 24 Microtubules in General Microtubules are long, hollow cylinders composed of tubulin heterodimers arranged in a specific fashion. Microtubules have an outer diameter of 25nm and are generally linear structures that are much more rigid than actin filaments. In most cultured cells microtubules are attached to a microtubule organising centre (MTOC) called a centrosome (Mitchison, 1984). Tubulin The tubulin heterodimer, 8 nm in length and approximately 100 kDa, is composed of two similar globular polypeptides called a-tubulin and p-tubulin, which combine to form microtubules. There are at least six different isoforms of both a-tubulin and p-tubulin and each is encoded by a different gene. Because the different forms of tubulin are very similar they will ordinarily co-polymerise into microtubules (Mitchison, 1984). Microtubule Structure The geometrical organisation of microtubules is achieved by the parallel arrangement of 13 long and linear protofilaments of tubulin surrounding a central hollow core. The protofilaments are composed of alternating a-tubulin and p-tubulin heterodimeric subunits stacked linearly end-to-end. Because the subunits are polar structures, the uniform arrangement of the heterodimer head to tail in the 13 protofilaments generates an overall polarity to the microtubule (Mitchison, 1984). 25 MAPS Microtubules are stabilised by microtubule-associated proteins (MAPs). MAPs provide microtubules with stability since they impede their disassembly. Since MAPs have been shown to bind to the sides of microtubules it is thought that they may provide a link between the microtubule and other cellular structures. High-molecular weight MAPs, which have molecular weights on the order of 200,000, include MAP 1 and MAP 2. Tau proteins, also members of the MAP family, have molecular weights on the order of 60,000. Motor proteins belong to the MAP family (Drechsel, et al., 1992). Microtubules in Sertoli Cells Microtubules are a very conspicuous component of the Sertoli cell cytoplasm. They are concentrated at the supranuclear region of Sertoli cells and are associated with apical crypts that surround developing spermatids. Most of the microtubules are aligned relatively parallel to the long axis of the Sertoli cells. The organization center of microtubules (MTOC) in Sertoli cells has not been identified. Microtubules also have been observed in the more basal regions (perinuclear regions) of Sertoli cells, however, their arrangement is less organised (Russell et al., 1993). Polarity of Microtubules in Sertoli Cells Microtubules surrounding developing spermatids are oriented such that slow growing or minus ends are situated apically, whereas their fast growing or plus ends are located basally (Redenbach and Vogl, 1991). Unlike in many other cells, the minus ends of Sertoli cell microtubules are not associated with 26 the perinuclear centrosome. In fact, in other cell types, such as neurons, the fast growing ends of microtubules are directed away from the soma (Heidemann et al, 1981). The Interaction of Microtubules and Motor Proteins Microtubules and actin filaments are polar structures. The polarity of these structures is important with respect to the direction of movement of enzymatic motors. For instance, myosins usually move towards the plus-end (barbed end) of actin fibers. Recently, however, myosin VI has been shown to move towards the minus-end of actin (Wells A.L., 1999). There are two main classes of microtubule motors that travel in opposite directions along microtubule tracts. Specifically, kinesins are generally but not exclusively plus-end directed microtubule motors whereas, dyneins are minus-end directed microtubule motors. Thus the polarity of microtubules determines the direction of cargo transport. In a motility assay if the polarity of the microtubule is defined then it is possible to predict the class of motor that is generating the observed microtubule movement (Kreis, 1993). Motor proteins are large molecules that appear, ultrastructurally, as elongate structures (40-100 nm in length) that have one or more globular heads connected to rod-like domains at their terminal ends. Kinesin and dynein have a tail domain with a characteristic structure. Other proteins may also form part of the motor and in the case of ciliary outer arm dynein, as many as ten of these associated proteins exist. Each of the 'main' domains (head, rod, tail) of motor proteins have specific functions. For instance, the S1 domain 27 of conventional myosin, contains actin-activated ATPase which generates force. In the case of kinesin, the globular domain of the heavy chain, contains an ATPase, a microtubule binding site and has motor activity. Within motor families the motor domains are usually highly conserved, however, different families of motors may show no sequence homology to one another. This diversity amongst the motor domains of the various families of motors suggests that motors either arose independently in evolution or very distantly in evolution (Kreis, 1993). In order for a motor to generate force one of the three following associations between the motor and the cytoskeletal elements must occur: 1. Directional transport is achieved when the motor binds to a cargo and then transports it along a cytoskeletal fibre. This mode of transport is widely used in the cytoplasm to transport elements such as membranous organelles, chromosomes and protein complexes. In this case, the attachment of the motor to the cargo must be highly specific to produce any useful activity and also to prevent motors from binding and tugging at structures at random. 2. Motors may also be used to generate a directional sliding force between parallel cytoskeletal filaments or fibres. Chromosome movement in anaphase B is accomplished using this type of motor/ cytoskeletal arrangement. 3. The third type of force generation is achieved when the motors self-assemble into a filament array and this filament array interacts with 28 adjacent substrate filaments. Muscle myosin filaments produce force in this way by moving on adjacent actin filaments. Unlike the mechanical or head domain of the motors, the tail end of the motors vary considerably in structure. This diversity may contribute to the way in which the motor is able to target a specific cargo. Motor Proteins - In General The microtubule-based spermatid translocation hypothesis indicates that enzymatic motors are responsible for the observed movement of spermatids during maturation. In the next few sections background information with respect to enzymatic motors is presented. Myosin was the first motor protein to be discovered by Kuhne in 1864, and it remained the sole motor under study, until dynein was isolated from tetrahymena cilia by Ian Gibbons in the 1960's. Muscle myosin and axonemal dynein became models for actin and microtubule based movements. Initially it was thought that these motors could produce many types of cytoplasmic motility. It is now known that there are many varieties of enzymatic motors and that they are often quite different from conventional muscle myosin or flagellar dynein (Kreis, 1993). In vitro motility assays, coupled with biochemical fractionation, led to the discovery of kinesin (Vale, 1987) and cytoplasmic dynein (Vallee et al., 1988). Once it was established that there was sequence homology present within motor families, molecular biological approaches were used to identify new motors within large motor superfamilies. Although, the degree of sequence 29 homology of the motor domains between kinesins and dyneins is generally 30-40%, the ATP binding domain is the most highly conserved region. In contrast to the motor domains, other elements of the motor show very little homology. The function of these non-motor regions has yet to be clearly defined; however, it has been suggested that these areas are involved with the targeting of motors to different macromolecular structures within the cytoplasm. Non-motor domains may be important in the regulation of functional activity of a particular motor. For instance, phosphorylation of the tail of myosin smooth muscle causes it to bend which inevitably results in an inhibition of the ATPase activity at the motor head (Kreis, 1993). Why are there so many different types of motor proteins? It has been proposed that cells probably target specific motors to different tasks (Kreis, 1993). Each individual motor may be unique due to the nature of its force generating ability or its direction of movement. There is, however, considerable overlap in the activities of most known motors. Thus, redundancy, which is especially critical to cellular activities such as cell division, is the most likely reason for having so many different motor types in the cytoplasm (Muresan et al., 1996). Notwithstanding the above, the myosin II motor is absolutely required to carry out cytokinesis. Thus, not every motor-requiring activity may be performed by several different motors (Kreis, 1993; Vallee et al., 1996). Brief History of Dynein Motors Cytoplasmic dynein was first described in 1987 by Paschal and colleagues. Gibbons and Rowe, however, first extracted the axonemal type of 30 dynein from the cilia of tetrahymena in 1965 and subsequently named it dynein: dyne=force, in= protein (Gibbons et al., 1965). This work by Gibbons led to the widespread acceptance of axonemal dynein as the motor protein responsible for flagellar and ciliary movements (Ogawa et al., 1996). In the 1970's when tubulin was found to be highly concentrated in cilia and flagella it was proposed that perhaps axonemal dynein interacted with polymerised microtubules and acted as an enzymatic motor. Later when it was determined that there was retrograde movement in addition to the anterograde movement of vesicles, produced by kinesin, the search for such a cytoplasmic motor was underway. This type of retrograde motor activity was first seen in nerve terminals. Eventually, microtubule associated protein 1C (MAP 1C) was extracted from rat brain (Paschal et al., 1987). MAP 1C was later re-named cytoplasmic dynein because there are similarities between this protein and the originally discovered axonemal dynein. For instance, both axonemal and cytoplasmic dynein motor proteins are microtubule activated ATPases (Ogawa et al., 1996). Cytoplasmic Dynein Introduction to the Role of Cytoplasmic Dynein Several studies have been conducted that examined the role of cytoplasmic dynein in microtubule-based minus-end directed organelle transport. In Schroer's (Schroer et al., 1989) study a reconstituted organelle motility assay was developed to test this hypothesis. They found that the movement of chick embryo fibroblast organelles on microtubules is driven by factors present in the cytosol. They then showed that cytoplasmic dynein was 31 present in the cytosol. When they inactivated dynein by vanadate-mediated UV photocleavage (photocleavage is specific for dynein heavy chain), they observed a marked reduction in minus-end directed organelle movement. Furthermore, when purified cytoplasmic dynein is added back to the UV-photocleaved cytosol the minus-end directed organelle movement resumes. Based on these observations they proposed, in 1989, that cytoplasmic dynein is a motor for minus-end directed organelle movement (Shroer et al., 1989). Membrane biogenesis and recycling require the movement of membraneous organelles, some of which depend on minus-end directed microtubule-based transport. In the 1970's it was suggested that 'saltatory' movements of endocytic organelles toward lysosomes are microtubule dependent (Freed et al., 1970). It is also thought that organelles such as the Golgi apparatus and lysosomes, which are located at the center of the cell during interphase, are actually retained there by an active mechanism (Schroer et al., 1989). Interestingly, when these cells are treated with microtubule polymerizing inhibitors the organelles disperse, however, once the microtubule structure is regained organelles are re-located towards the centrosome. The direction of organelle movement in this scenario would be consistent with a minus-end directed motor. Schroer predicted, based on their findings, that receptors located on organelles would be specific for and recruit minus-end directed motors, such as cytoplasmic dynein, and thus, provide a mechanism to control transit to the minus-end of the microtubule. Therefore, there is 32 evidence for organelle transport mediated by minus-end directed microtubule motors. Structure of Dynein Cytoplasmic dynein is an extremely complex and large protein that is a member of the dynein superfamily of proteins. Dynein is composed of two and sometimes even three heavy chains which are large globular force-generating heads that have a total molecular weight of 1.2-2 x 106 kDa (Figure 13). Cytoplasmic dynein has two such globular heads of 530 kD each (4644 amino acids in rat) which fold to form phosphate binding P-loops at its central core. Each dynein motor also has several subunits associated with it, which are consistent with each of the four polypeptide classes: the heavy, intermediate, light intermediate and light chains. The ATPase activity occurs in the heavy chain of the molecule, which is the force producing area. As many as fifteen dynein heavy chain genes are present in each of the organisms examined to date (Vallee et al., 1998). Both axonemal and cytoplasmic dynein are similar in size and exhibit considerable homology along their length. This finding is unlike that for the myosin and kinesin families of motor proteins where numerous and different subfamilies have been identified. Dynein Heavy Chain The dynein heavy chain or motor domain is more complex than those of other motor proteins. Each dynein heavy chain is composed of four P-loop sequence elements, which are the nucleotide binding and hydrolysis areas and are located at the C-terminus of the molecule. The P-loop elements are 33 separated by 35-40 kDa intervals within the central area of the molecule. Mutations of the P-loop element of the dynein molecule prevent the molecule from producing any force (Vallee et al., 1998). Other Elements of the Dynein Motor Protein Dynein is composed of three intermediate chains (74 kD each) and four light intermediate chains (55 to 60 kD each). Since, cytoplasmic dynein must simultaneously associate with microtubules and cargo, it has been suggested that one end of the cytoplasmic dynein motor (COOH terminal) invariably interacts with microtubules while the other end (NH2 terminal) interacts with the cargo (Hirokawa, 1998). The cargo binding site of the dynein motor protein may be modulated and as such be specific for a particular type of cargo. It has been suggested that the dynactin complex serves this very purpose (Hirokawa, 1998; Shnapp et al., 1989; Shroer et al., 1989; Holzbauer et al., 1994). The Dynactin Complex Dynactin is composed of 10 subunits: p150 g l u e d (glued, since it is associated with the glued gene) or p135 g l u e d, p62, dynamitin, actin-related protein 1 (Arp 1), actin, actin capping protein a subunit, p27 and p24. The p150 g l u e d and p135 g l u e d heterodimers project from the Arp 1 component of the dynactin complex. The NH2-terminal of p150g l u e d forms a side projection that interacts with the 74kD intermediate chain. When p150 g l u e d binds to cargo it is linked to the heavy chain of the motor complex via the 74kD intermediate chain which is linked to the Dynamitin region via the 55kD light intermediate chain. p150 g l u e d is also probably linked to the Arp 1 and dynamitin sections of the 34 dynactin complex directly. Interestingly, p150 g l u e a can also bind to microtubules directly. It is thought that the dynactin complex may be the receptor for dynein on membranous organelles (Hirokawa, 1998; Gill et al., 1991). Microtubule-Binding Stalks A microtubule binding site at the heavy chain of cytoplasmic dynein has been identified. An examination of cytoplasmic dynein molecules lacking the entire P-loop which were in the presence of microtubules revealed that motors and microtubules colocalized. Microtubule binding activity was mapped to a site approximately 340 amino acids residues downstream from the fourth P-loop element. It has been proposed that at the end of the large cytoplasmic dynein head resides a stalk which binds to microtubules. The stalk is predicted to be composed of a small microtubule-binding element (residues 3276-3407) which is flanked on either end by a similar region of about 100 residues that form coiled-coil a helices. It is predicted that these two coiled coils associate with each other and form an antiparallel coiled-coil. This structure would thus, have at its distal end a microtubule binding domain and a stalk of approximately 15 nm in length. This complete structure is thought to be positioned at the tip of the cytoplasmic dynein head (Vallee et al., 1998). It is thought that the role of the stalk element of the cytoplasmic dynein is to allow for several dynein molecules to bind to the same microtubule. It would be impractical for several dynein heads, as large as they are, to associate with adjacent tubulin monomers of the same microtubule. However, if a stalk, of 35 approximately 15 nm, was positioned at the tip of each cytoplasmic dynein head then the interaction between adjacent cytoplasmic dyneins with adjacent tubulin monomers of the same microtubule becomes much more likely. It is also important to consider the length of the proposed dynein stalk since if the stalk is too short then efficient dynein motor activity would be less likely, especially if on the same microtubule; however, if the stalk was too long then the force produced by the dynein head would not be transferred effectively to the microtubule. How is the energy generated from ATP hydrolysis able to produce a locomotive force if the contact between the cytoplasmic dynein head and the microtubule is via a stalk? There are at least two possible mechanisms that could account for such an energy transfer. Firstly, the stalk could act as a lever arm that amplifies conformational changes occurring in the cytoplasmic dynein head, where ATP hydrolysis occurs. Secondly, the stalk may serve as a passive arm that the dynein head forcefully moves as a result of a conformational change that occurs at the globular head structure. Based on images of dynein motors associated with axonemes it has been suggested that a large change in the overall length of the dynein molecule, due to elongation and contraction of the molecule, is occuring during ATP hydrolysis. This information would support either of the two mechanisms suggested for force transmission through the stalk (Vallee et al, 1998). 36 Control of Dynein Mechanics How does dynein mediate movement along a microtubule? In the absence of ATP, dynein binds to microtubules, in a fixed position. When ATP becomes available for binding the dynein stalk dissociates from the microtubule. Since the contact between the dynein head and the microtubule is through the stalk's terminal end point and because the stalk has been shown to preserve its general morphological structure during the complete crossbridge cycle, it is unlikely that ATP binding events trigger major conformational changes within the dynein stalk. However, it is possible that minor changes within the dynein stalk tip do occur, because of communication between the dynein head and stalk tip, which could explain the attachment and subsequent detachment of the stalk tip from the microtubule. Crossbridging between the stalk tip and the microtubule may also be due to a temporary change in the orientation (such as rotation) between the stalk and microtubule. When the stalk tip binds the microtubule, prior to ATP hydrolysis in the dynein head, a signal must be transmitted to the dynein head so that it undergoes a power stroke. Once the power stroke peaks, the dynein stalk tip is released from the microtubule so that it can rebind to the microtubule at a new and progressed position. The cycle would then begin again with the cytoplasmic dynein head initially binding ATP for hydrolysis (Vallee et al., 1998). Cytoplasmic Linker Proteins (CLIPs) The movement of cytoplasmic organelles is dependent on effective interaction between microtubules and motor proteins. The organelles or cargo, 3 7 to be transported, must be appropriately linked to the motors which must in turn interact with microtubules (Kreis 1993). Binding assays to study the association between microtubules and organelles have been established and two proteins, namely: CLIP-170 and a similar protein of 50 kDa, exist. The structure of CLIP-170 has been deduced from its amino acid sequence, however, the structure of the 50 kDa protein is still unknown (Kreis 1993; Vaughan et al., 1999). CLIP-170 CLIP-170 is an elongated (2.5nm by 110 nm), homodimeric molecule. Its structure resembles that of myosin and kinesin. CLIP-170 does not contain the known nucleotide binding sites that are generally found in microtubule motors, and thus it is unlikely that it is a motor. However, a tandemly repeated motif has been identified in its N-terminal domain which is involved in microtubule binding and one of these repeats is also present in other proteins such as dynactin (Kreis 1993; Vaughan et al., 1999). CLIPs may be an important factor in the initiation of organelle movement especially along microtubules. CLIPs may form an initial contact between vesicles or organelles and the microtubules and in so doing stabilise the cargo while the motor 'picks-up' the cargo. Once an active motor has bound its cargo then the link (CLIPs) between vesicles or organelles and microtubules must be weakened. It has been shown that once CLIP-170 binds to microtubules it then becomes sensitive to phosphorylation by a kinase present in the microtubule binding fraction. Thus, phosphorylation of CLIP-170 may be the 38 'switch' that allows the CLIP-170 to release from the organelle and the microtubule. 39 Microtubules associate with ER It is well-known that microtubules can closely associate with ER. In 1971, Franke showed, using EM, that microtubules can be closely associated with adjacent ER. In fact, he identified crossbridges linking the ER to microtubules in cells. In 1986 Terasaki used fluorescent labelling techniques to study the ER and microtubules in fixed cells. Interestingly, when the ER was stained with 3,3'-dihexylocarbocyanine Iodide (DiOC6) it appeared as a polygonal network which was associated with microtubules, especially in the lamellipodia where individual microtubules and the ER tubules could be observed. In addition, he noted that when the microtubules in the cells were depolymerized, the ER network gradually retracted toward the cell center. Furthermore, if the microtubules repolymerized the ER network extended to its original organisation. From this study he concluded that microtubules play an important role in maintaining the structure of the ER (Teresaki, 1986). In 1998 a study by Klopfenstein showed that a 63 kDa protein which is a non-glycosylated type II integral ER membrane protein could actually link microtubules to the ER directly (Klopfenstein et al., 1998). Studies in which microtubule-depolymerization drugs were used also indicated that microtubules and the ER are closely related. For example in a study by Vogl et al. (1983) where depolymerization drugs were used in intact Sertoli cells of the ground squirrel it became clear that the state of the microtubules can greatly influence the organization of the related ER. 40 Based on EM studies it has been shown that the ER has a large surface area and that it often associates with microtubules (Dabora et al, 1988). Dabora showed that a network similar to an ER can form (in vitro) from membranous aggregates in the presence of microtubules and ATP. In their study they proposed a mechanism for the formation of membranous in vitro networks, in which, cisterns of membranes are formed from membrane aggregates by the translocation of point contacts along microtubules. Dabora referred to this process as 'microtubule-dependent' tethering. This model proposes that ER can move along stationary microtubules (Dabora et al, 1988). Cytoplasmic Dynein and Apically - Directed Spermatid Translocation Cytoplasmic dynein is a minus-end directed microtubule dependent enzymatic motor. Cytoplasmic dynein could thus, due to the orientation of Sertoli cell microtubules and their proximity to the ER of ES, be responsible for the apical translocation of developing spermatids (Figures 3, 5 and 6). In support of this prediction is a study that showed that the intermediate chain of cytoplasmic dynein (IC74) is localised to the area surrounding developing spermatids (Miller et al., 1999). Interestingly, a minus-end directed kinesin motor has been identified in Drosophila called Ned, which is required for normal meiosis and differs from axonal kinesin both in the direction and rate at which it moves along microtubules (Alberts et al., 1994). Thus it is theoretically possible that a spermatid-specific minus-end kinesin motor could be responsible for this translocation. 41 Spermatid translocation occurs in both the apical and basal directions; however, there is no single enzymatic motor capable of moving to both the plus- and minus-ends of microtubules. Thus, I predict that cytoplasmic dynein is most likely the motor involved with apically directed spermatid translocation and that a plus-end directed microtubule motor, such as kinesin, may be responsible for the basally directed spermatid translocation event. 42 Figure 5. - Structural Features of Ectoplasmic Specializations and the Spermatid Translocation Model. Panel A: A high-magnification EM of a cross section through the head of an elongate spermatid. This micrograph reveals the important structural relationships between the ES junctional plaque and the spermatid plasma membrane. Magnification: X 72,000, bar= .25 um. Panel B: A schematic figure that depicts the translocation hypothesis. The polarity of the microtubules, as indicated, is important with respect to the selection of enzymatic motors and the translocation process. Panel C: A high-magnification EM of a longitudinal section through the ES junctional plaque. The ER cisterns of the ES are clearly visible. Magnification: X 69,000, bar= .25 um. 43 Brief Introduction to the Motility Hypothesis It is hypothesised that the ES, together with its spermatid, move, as a single unit, parallel to the long axis of the Sertoli cell, first basally (Figure 6, panel 1 to 2) and then apically (Figure 6, panel 2 to 3) along microtubule tracts surrounding an elongate spermatid (see microtubules - Figure 3). The ES plaque is then disassembled prior to spermiation, and the mature spermatid is released to the lumen of the tubule (Figure 6, panel 4). The structure of the ES, particularly the fact that the ER is coupled, via a layer of actin filaments, to the plasma membrane of the Sertoli cell strongly supports the prediction that the ES moves as a cohesive unit (Vogl et al. 1991). • It is predicted that motor proteins, such as dynein and kinesin are anchored to the cytoplasmic face of the ER (labelled as motor proteins on Figure 5, middle panel, see also Figure 13 for relationship between dynein and ER) and move along adjacent microtubules and thus, mediate the translocation of spermatids. 45 Figure 6. - The Spermatid Translocation Model A four panel schematic diagram (panel one is on the left) illustrating the spermatid translocation process, taken from Vogl et al., 1991. Note that the spermatid is first shuttled to the basal region of the Sertoli cell (first panel to second panel), then back to the apical region of the Sertoli cell (second panel to third panel) and is then released to the lumen of the seminiferous tubule. The apical ES plaque disassembles prior to spermiation whereas the basal ES plaque disassembles to allow maturing spermatogonia to move apically from the basal compartment to the adluminal compartment. 46 The Thesis Problem Hypothesis The hypothesis to be tested in this study is that cytoplasmic dynein motors are anchored to the cytoplasmic face of the ER component of ES. Approach To test my hypothesis I used a morphological technique, electron microscopy and a biochemical technique, SDS-PAGE and immunoblotting. Immunoelectron microscopy was used to verify that antibodies, raised in a mouse and specific to the intermediate chain of the dynein motor complex (IC74), react positively on the cytoplasmic face of the ER component of the ES (Chapter Two). Western blot analysis was used to show that antibodies, raised in a rabbit and specific to the heavy chain of the dynein motor complex (DHC1a) (and IC74 as discussed above), reacted positively with proteins on immunoblots and that they are visible as distinct bands at the appropriate molecular weights when tested on supernatants from mechanically fragmented and gelsolin treated rat spermatids. Gelsolin is an actin severing enzyme and thus, in this system gelsolin provides a method of enriching for the ER component of the ES junctional plaque (see Gelsolin Experiment, Figure 17). Rationale Spermatids are translocated to the apical region of Sertoli cells during their maturation. We predict that minus-end directed microtubule dependent 48 dynein motors are anchored to the cytoplasmic face of ER and are responsible for this apically-directed translocation event. The rationale for the use of the dynein intermediate chain antibody (IC74-monoclonal) is twofold. Firstly, this antibody (BAbCo, Richmond, CA) was previously used in our laboratory in a florescence microscopy study on the same tissue (Miller et al., 1999). That study clearly indicated that the IC74 component of the dynein motor complex is present around the spermatid head. This present study, is aimed at more precisely localising cytoplasmic dynein at the level of the ES. Secondly, the IC74 intermediate chain specific antibody is widely used and is available commercially (I also tested a sample of the original anti-IC74 produced in Dr. Pfister's laboratory). The rationale for using the heavy chain specific antibody in our system is also twofold. Firstly, previous studies have indicated that the DHC1a is present in testes and is associated with the IC74 component of cytoplasmic dynein (Criswell et al., 1998). Secondly, based on previous studies, the heavy chain component of cytoplasmic dynein motor is a non-variable element of the cytoplasmic dynein motor complex. The presence of the dynein HC1a in our tissue would provide strong evidence in support of our prediction that a cytoplasmic dynein motor is localised to the cytoplasmic face of the ER component of ES. 49 CHAPTER2 CYTOPLASMIC DYNEIN IS LOCALIZED TO THE ER COMPONENT OF ES: MORPHOLOGICAL EVIDENCE Introduction Based on motility studies currently underway in our laboratory and on previous work it became important to determine the precise location of the cytoplasmic dynein motor protein at the level of the ES (Beach et al.,1999 and Miller et al.,1999). In this Chapter I present and discuss the morphological data that I collected using standard immuno-EM protocols. I used two protocols, namely: 1. The pre-embedding EPON labelling process. 2. The post-embedding Lowicryl and Unicryl labelling processes. The EM data presented in this Chapter is consistent with results obtained previously at the light microscopy level (Miller et al., 1999) and with the spermatid translocation model. Based on the spermatid translocation model I propose that the cytoplasmic face of the ER component of the ES is the most likely anchoring site for cytoplasmic dynein motors. The results from my EM study strongly support this hypothesis. 51 Materials and Methods Animals All animals used in this study were reproductively active male Sprague-Dawley rats with an average weight of approximately 400g. They were obtained from a colony maintained at the Animal Care Facility at the University of British Columbia. The animals were cared for and used in accordance with guidelines established by the Canadian Council on Animal Care. Reagents and Antibodies All chemicals and reagents used in this study were purchased from the Sigma-Aldrich Company (St. Louis, MO, USA) or the Fisher Scientific Co. (Suwanee, GA, USA), unless otherwise noted. Glutaraldehyde, sodium cacodylate, osmium tetroxide, Epon 812, Lowicryl and Unicryl were obtained from J.B. EM services (Dorval, Quebec). The antibody specific to the IC74 component of cytoplasmic dynein (Dillman J.F. et al., 1994), was obtained from either the BAbCo Co. (Richmond, CA, USA) or as a gift from Dr. Pfister (Univ. of Virginia Health Sciences Center, Charlottesville, USA). The antibody specific to the heavy chain of cytoplasmic dynein (DHC1a) was received as a gift from Dr. Asai (Purdue University, West Lafayette, IN, USA). The monoclonal antibody to Glucose Regulated Protein (GRP94) was purchased form Stressgen Biotechnologies Corp. (Victoria, BC). The secondary gold-conjugated (10 nm) antibody was purchased from Jackson ImmunoResearch Laboratories Inc. (West Grove, PA, USA). The secondary horse-radish peroxidase (HRP) antibodies were purchased from 52 Santa Cruz Biotechnology Inc., (Santa Cruz, CA, USA). The molecular weight markers (Kaleidoscope Prestained Standards), acrylamide and TRIS-HCI were purchased from Bio-Rad Laboratories (Hercules, CA, USA). Orchotomy Rats were placed in a chamber where they were anaesthetised with halothane (setting 3.5/5, delivered with 0 2 ) . Once the animals were under deep anaesthesia (this ordinarily required six to eight minutes) they were placed on a surgical table where they were fitted with a mask that continued the delivery of halothane. After positively confirming, by pinching the rats' rear paws with hemostats, that their withdrawal reflexes were absent, the surgical removal of their testis was performed. The testis were removed from the scrotum and the epididymis was cut away. The spermatic cord was clamped and cut near its entry point at the pole of the testis. The organ was then processed for either immuno-EM or western analysis. The rat was euthanized by cutting open the thorax and by severing the heart and lungs with a scissor. 53 Electron Microscopy- Pre-embedded Labelling For the Immuno-EM study, aimed at precisely localizing cytoplasmic dynein motors, at the level of the ES, an anti-dynein antibody (ant-IC74) was used. Testes were removed from anaesthetised animals as described above. Organs were perfused briefly (2 minutes), via the testicular artery, firstly with phosphate buffered saline (PBS: 150 mM NaCI, 4.0 mM Na/K P0 4, 5.0 mM K, pH 7.3), to clear blood from vascular system, and then with fixative (30 minutes) (PBS solution containing 3.0% paraformaldehyde at 33 ° C). Following fixation, the testis were perfusion washed with PBS for 30 minutes, cut into small fragments with razor blades and then further fragmented by aspirating the tissue through a needle (18 G) mounted on a 1 cc syringe. The fragmented tissue was then allowed to settle through approximately 10 mis of PBS for 20 minutes. The upper third of this material was then collected for immunostaining and then divided into four equal aliquots, as follows: (1) Experimental sample, (2) IgG control sample, (3) Secondary antibody control sample, (4) Blank sample. Samples were then pelleted by centrifugation (setting 6 for 4 minutes on an Eppendorf table top centrifuge) and then re-suspended in blocking solution (PBS/ 0.1% bovine serum albumin (BSA)/ 5% normal goat serum (NGS)/ 0.05% Tween 20) for 10 minutes at room temperature. Experimental samples were then re-suspended in primary antibody solution: monoclonal anti-IC74 at a concentration of 1:25 (BAbCo, Richmond, USA or from Dr. Pfister, Univ. of 54 Virgnia-HSC, USA), for 120 minutes at room temperature. Controls were incubated in normal mouse IgG (diluted to the same protein concentration as the experimental samples) and under otherwise identical conditions. Secondary antibody controls and double blank controls were incubated in PBS/ 0.01% BSA/ 0.05% Tween 20 and otherwise identical conditions. Samples were then washed with PBS, containing 0.1% BSA and 0.05% Tween 20, three times. All samples, except for the double blank controls were then incubated in gold-conjugated secondary antibody for 120 minutes at room temperature and at a concentration of 1:50. The double blank controls were incubated in PBS/ 0.01% BSA/ 0.05% Tween 20. Samples were then washed, as described above, pelleted by centrifugation, and then fixed with PBS containing 3% paraformaldehyde and 0.5% glutaraldehyde (pH 7.3). The pellets were then washed with 0.1 M sodium cacodylate (pH 7.3), post-fixed on ice for 1 hour in 1% Osmium Tetroxide diluted in 0.1 M sodium cacodylate (pH 7.3), washed with ddH 20, and then stained en bloc in 1% uranyl acetate in ddH20. Following a final series of washes the samples were dehydrated through a series of ethyl alcohol solutions (30%, 50%, 60%, 70%, 90%, 95%, 100%) and embedded in Epon. Serial sections (70 nm) were cut, using an ultramicrotome fitted with a 2 mm diamond knife, and collected on 200-300 mesh copper grids. Some sections were then stained with uranyl acetate and lead citrate and all were evaluated for specific immunoreactivity with a Philips 300 electron microscope operated at 60kV. 55 Repetition of Experiments When I initially began probing the Epon embedded tissue with antibodies specific to the IC74 (obtained from BAbCo) component of dynein I observed, early on, some very encouraging results. However, unaware that not all anti-IC74 monoclonal antibodies are equally sensitive, I experimented with a less expensive anti-IC74 antibody from Chemicon International Inc. (Temecula, CA). After several trials it became clear that the data collected with this antibody was not consistent with my earlier. I repeated the experiment using the non-reactive antibody twelve times. Subsequent to this I re-tried the anti-IC74 antibody available from BAbCo and positive results resumed. I also obtained a sample of the anti-IC74 antibody from Dr. Pfister, who originally developed this particular antibody (Dillman J.F. 3 r d et al., 1994). The 'Pfister' anti-IC74 antibody provided data that was indistinguishable from the 'BAbCo' anti-IC74 study. I repeated the 'BAbCo' anti-IC74 experiment four times and the 'Pfister' anti-IC74 experiment was done once. In addition, western blot analysis confirmed that the anti-IC74 antibody was specific (Miller et al., 1999) and that it did not react with bovine brain tubulin. Electron Microscopy- Post-embedding Labelling In contrast to the pre-embedding immuno-EM protocol, described above, this protocol, referred to as post-embedding immuno-EM, identifies specific immunoreactivity on whole sections of seminiferous epithelium cut from fixed rat testis embedded in Lowicryl or Unicryl resin. 5 6 Testis were obtained, as described above, perfused with PBS (2-3 minutes) and then with a fixative containing 3% paraformaldehyde/ 20mM ethylacetimidate/ PBS (pH 7.3), for 30 minutes, followed by perfusion with a second fixative containing 3% paraformaldehyde and 0.1% glutaraldehyde in PBS for 30 minutes. Testis were then perfusion washed with PBS for 30 minutes, cut into small cubes, immersion washed in 50mM NH4CI in PBS for 30 minutes and then further washed by immersion in only PBS for 30 minutes. The tissue cubes were then dehydrated in a series of methanol solutions (30%, 50%, 70%, 90%, 95%, 100%x3- 60 minutes each step). Tissue cubes were then embedded in Lowicryl or in Unicryl resin and polymerized at -20 °C using ultraviolet light. Cured blocks were stored at 4 °C. Cured blocks were sectioned using an ultra microtome fitted with a 2 mm diamond knife. Sections (70 nm) were collected on formvar/carbon coated 200-300 mesh nickel grids for immunolabelling. Grids were divided into four groups (experimental, IgG control, secondary control, double blank). All grids were initially placed on a drop of blocking buffer (PBS/ 0.01 M Glycine/ 0.1% BSA/ 1:20 NGS/ 0.05% Tween 20) for 10 minutes. Experimental grids were then placed on a 40 uJ drop of primary antibody (anti-IC74 at cone, of 1:10 diluted in PBS/ 0.1 M Glycine/ 0.1% BSA/ 1:20 NGS/ 0.05% Tween 20) for 120 minutes and at room temperature. IgG control grids were placed on a 40 ui drop of normal mouse IgG (diluted to the same protein concentration as experimental sections), whereas, secondary controls and double blank grids were placed on a 40 uJ drop of the diluting 57 solution (PBS/ 0.1 M Glycine/ 0.1% BSA/ 1:20 NGS/ 0.05% Tween 20) for a period of 120 minutes and at room temperature. All grids were then washed, as above, in PBS/ 0.01 M Glycine/ 0.1% BSA (3 washes, 3 minutes each wash). Experimentals, IgG controls and secondary control grids were then placed on a 40 JJ:I drop of 10nm gold- conjugated secondary goat anti-mouse IgG antibody (at a concentration of 1:25 made up in PBS/0.01% BSA/ 0.05% Tween 20) for 120 minutes and at room temperature. Double blank grids were placed on a 40 ul drop of PBS/0.01% BSA/ 0.05% Tween 20 for a period of 120 minutes and at room temperature. All grids were then washed twice, as described above, postfixed in 2.0% glutaraldehyde diluted in PBS (10 minutes) washed as indicated above, stained with aqueous 1% uranyl acetate in dH20 and then washed three times with dH 2 0 and allowed to air dry. Grids were then examined for immunoreactivity and compared with controls using a Philips 300 electron microscope operated at 60 kV. Post- Embedding vs. Pre-Embedding Despite the fact that the two embedding protocols (Epon or Unicryl and Lowicryl), are technically very different, they did provide similar immuno-EM results. For my studies, however, I believe that the pre-embedding labelling (Epon) technique is superior for the following three reasons, namely: 1. The Epon protocol requires the fragmentation of the tissue to separate the spermatid from the Sertoli cell. The ES is rarely ripped away from the spermatid head and in fact, it is the Sertoli cell plasma membrane that tears to allow the ES to remain attached to the spermatid head. This 58 fragmentation step exposes large regions of the cytoplasmic face of the ER (the predicted site of the cytoplasmic dynein motor proteins) to antibodies that are specific to the dynein motor complex. The post-embedding labelling procedure does not necessarily expose large areas of the cytoplasmic face of the ER for antibody binding. 2. The Epon technique excellently preserves, by means of post-labelling fixation with osmium tetroxide, the membranous structures of the spermatids. The post-embedding labelling technique (Lowicryl or Unicryl), which does not employ osmium tetroxide, poorly preserves the membraneous structures and thus the EM images of the membranous structures are less distinct as compared to the Epon protocol. 3. A disadvantage of the Epon technique is that even though the spermatids are mechanically stripped away from the Sertoli cell, the cytoplasmic face of the ER (where the dynein motors are predicted to be) can be obscured by Sertoli cell cytoplasm or by other cellular elements. Thus, the observed sensitivity of the IC74 antibody to dynein may under-represent the actual concentration of dynein motor proteins present at the cytoplasmic face of the ER. There are two advantages to the post-embedding protocol (Unicryl or Lowicryl). 1. The post-embedding labelling technique allows for a comparison of the presence of gold particles at the predicted antigen site (cytoplasmic face of ER) with other areas of the tissue section that are not predicted to be associated with cytoplasmic dynein. Thus, it is possible to compare the 59 presence of gold at the cytoplasmic face of the ER with areas that are not expected to present dynein proteins such as, on the nucleus of the spermatid or in the basal cytoplasm of the Sertoli cell. 2. When the Unicryl and Lowicryl resins are sectioned, the face of the section tends to be jagged (unlike Epon which presents a very smooth surface). This is because the section is actually 'cracked off from the block face. This jagged surface presents a large surface area which will come into contact with more of the antibody incubation solution. This feature of the resins may promote increased antigen exposure for these studies. 60 Results Electron Microscopy The anti-IC74 antibody (BAbCo only) was tested, using western analysis, on bovine brain tubulin to confirm that the antibody does not cross react with tubulin. The results from this study indicated that the antibody does not react with bovine brain tubulin. Pre-embedded immuno-gold study I predicted, based on the spermatid translocation model that, cytoplasmic dynein is localised to the cytoplasmic face of the ER component of ES. To test my hypothesis immuno-gold studies were performed. The fragmented tissue that was labelled prior to embedding showed excellent preservation of the structural elements forming the ES plaque. Thus, the identification of the ER component of ES, in these studies, was easily performed. Probing the fragmented tissue with the anti-IC74 dynein antibody and then examining these grids with high-magnification EM revealed that the great majority of visible gold particles were present, usually in 2-4 particle clusters, on or near the cytoplasmic face of the ER (Figure 7A, 7B). At higher magnification several gold particles were positioned on the cytoplasmic face of well defined cisterns of ER (Figure 8A). On cells which had an incomplete ER component of ES, any gold particles present, at that particular plane of section, were generally situated on the cytoplasmic face of a cistern of ER (Figures 9B, 9C). Although, gold particles in the majority of sections were highly concentrated to the cytoplasmic face of ER cisterns that remained anchored to 61 the actin layer of ES (Figure 8A, 9B), there were certainly some gold particles that were visible on other membraneous structures present in the Sertoli cell cytoplasm surrounding the apical crypts. The controls indicated that the immuno-reactivity is specific. Very few gold particles were present when the primary antibody was replaced with normal mouse IgG (Figures 7C, 8C). Figure 8D specifically demonstrates that on secondary controls with a complete and attached ER that there were no visible gold particles. Gold particles were also absent when the primary and secondary antibodies were replaced with buffer alone (data not shown). Post-embedded immuno-gold study In tissue that was labelled after embedding, the morphology of the ES plaques was not as good as compared with tissue labelled prior to embedding. Probing sections of seminiferous epithelium with the anti-IC74 dynein antibody and then examining the grids with high-magnification EM revealed linear clusters of gold particles lying on what I interpret to be, based on previous actin localization studies, the cytoplasmic face of the ER (Figures 10A, 10B, 11A-E). The head and non-ER regions of ES are relatively devoid of gold particles (Figure 10A, 11B, 11C, 11D). Similar to the results collected from the epon-embedded IC74 probed tissue, gold particles were also present in areas of the Sertoli cell cytoplasm slightly removed from direct contact with ESs (Figures 10B). Also, consistent with previous experiments, (Hall E.S., 1992; Miller et al., 1999; Paschal et al., 1992) there was a strong presence of IC74 staining in the tail (Figure 12B) and manchette regions (Figure 12A) of 62 spermatogenic cells. This staining was specific and absent from the more general cytoplasmic areas of these cells. The controls indicate that the immuno-reactivity is specific. Gold particles were not present when the primary antibody was replaced with normal mouse IgG (Figure 10C), nor were there gold particles present when the primary antibody was replaced with buffer alone (data not shown). 6 3 Figure 7. - Pre-embedded Immuno-gold Study Targeting the IC74 Component of Cytoplasmic Dynein at the Cytoplasmic Face of the ER Component of ES. Shown here are electron microscopic images of late-stage spermatids collected from mechanically fragmented testis. This tissue was processed for standard EM and was embedded in EPON. These spermatids have an intact ES. The cytoplasmic face of the ER is indicated in panel D. Panel A. A typical high magnification micrograph of a spermatid incubated with an anti-dynein (IC74-raised in mouse) monoclonal (BAbCo) antibody followed by incubation with an anti-mouse 10 nm gold conjugate. Gold dots (arrows) indicate the presence of the IC74 component of the cytoplasmic dynein motors in association with the cytoplasmic face of the ER. Magnification: X 38,000, bar = 0.5 um. Panel B. This micrograph is of a late-stage spermatid, processed in the same manner as the one shown in panel A. Clearly, visible are the plasma membranes of both the Sertoli cell and spermatid and multiple cisterna of ER. The Gold dots (arrows) are indicative of the location of IC74 component of cytoplasmic dynein motors adjacent to the cytoplasmic face of ER. Magnification: X 34,900, bar = 0.5 um. Panel C. This micrograph of a late-stage spermatid, incubated with normal mouse IgG is devoid of gold dots. Clearly visible is the ER component of the ES. Magnification: X 40,400, bar = 0.5 um. Panel D. This micrograph is of a late-stage spermatid, processed in the same manner as the one shown in panel C. Magnification: X 37,250, bar = 0.5 um. 64 Figure 8. - Pre-embedded Immuno-gold Study Targeting the IC74 Component of Cytoplasmic Dynein at the Cytoplasmic Face of the ER Component of ES. Shown here are electron microscopic images of late-stage spermatids collected from mechanically fragmented testis. This tissue was processed for standard EM and was embedded in EPON. These spermatids have intact ESs. The cytoplasmic face of the ER is indicated in panel D. Panel A. This is a typical high magnifieatipn micrograph of a spermatid incubated with an anti-dynein (IC74- mouse) monoclonal (Pfister) antibody followed by incubation with an anti-mouse 10 nm gold conjugate. Gold dots (arrows) indicate the ER- associated IC74 component of the cytoplasmic dynein motors. Magnification: X 56,200, bar = 0.5 um. Panel B. This micrograph is typical of a late-stage spermatid, processed in the same manner as the one shown in panel A, but at higher magnification. Clearly visible are the cisterna of ER. The Gold dots (arrows) indicate the location of the IC74 component of cytoplasmic dynein motors adjacent to the cytoplasmic face of ER. Magnification: X 68,100, bar = 0.5 um. Panel C. This typical micrograph of a late-stage spermatid, incubated with normal mouse IgG is devoid of gold dots. Clearly visible is the cytoplasmic face of the ER component of the ES. Magnification: X 40,400, bar = 0.5 um. Panel D. This typical micrograph is of a late-stage spermatid, incubated solely with an anti-mouse 10 nm gold conjugate. Clearly visible are discrete cisterna of ER. Magnification: X 35,200, bar = 0.5 um. 66 Figure 9. - Pre-embedded Immuno-gold Study Targeting the IC74 Component of Cytoplasmic Dynein at the Cytoplasmic Face of the ER Component of ES. Shown here are electron microscopic images of late-stage spermatids collected from mechanically fragmented testis. This tissue was processed for standard EM and was embedded in EPON. These spermatids have intact ESs. The cytoplasmic face of ER is indicated in panel D. Panel A. High magnification micrograph of a spermatid incubated with an anti-dynein (IC74- mouse) monoclonal (BAbCo) antibody followed by incubation with an anti-mouse 10 nm gold conjugate. Gold dots (arrows) indicate, the presence of the ER- associated IC74 component of the cytoplasmic dynein motors. Magnification: X 67,000, bar = 0.5 um. Panel B. This micrograph is of a late-stage spermatid, processed in the same manner as the one shown in panel A, but at lower magnification. The Gold dots (arrows) indicate the location of the IC74 component of cytoplasmic dynein motors which are along the cytoplasmic face of the ER membrane. Magnification: X 23,300, bar = 0.5 um. Panel C. This micrograph is of a late-stage spermatid, processed in the same manner as the one shown in panel B, but at slightly higher magnification. The Gold dots (arrows) indicate the location of the IC74 component of cytoplasmic dynein motors associated with the ER. Magnification: X 34,760, bar = 0.5 um. Panel D. This micrograph of a late-stage spermatid, incubated with normal mouse IgG is devoid of gold dots. Clearly visible is the cytoplasmic face of the ER component of the ES. Magnification: X 42,600, bar = 0.5 um. Panel E. This micrograph is of a late-stage spermatid, processed in the same manner as the one shown in panel D. Magnification: X, bar = 0.5 um. 68 £ 1 Figure 10. - Post-embedded Immuno-gold Study Targeting the IC74 Component of Cytoplasmic Dynein at the Cytoplasmic Face of the ER Component of ES. Shown here are electron micrographs of late-stage spermatids that were processed for standard EM and embedded in Lowicryl. After ultra sectioning these grids were incubated with an anti-cytoplasmic dynein (IC74) antibody followed by an anti-mouse gold (10 nm) conjugate. Panel A. Shown here is a high magnification micrograph of a late-stage spermatid. Gold dots (especially those in the linear group (arrows at bottom right of figure) indicate the location of the IC74 component of cytoplasmic dynein. The gold is localized to the area of the ER membrane. Magnification: X 51,000, bar = 0.5 um. Panel B. Shown here is a micrograph similar to the one shown in panel A. The gold dots are located along the membrane that is consistent with the location of the cytoplasmic face of ER. Magnification: X 42,000, bar = 0.5 um. Panel C. Shown here is a high magnification micrograph of two late-stage spermatids. This grid was incubated with normal mouse IgG and is devoid of gold dots. Magnification: X 47,800, bar = 0.5 um. 70 Figure 11. - Post-embedded Immuno-gold Study Targeting the IC74 Component of Cytoplasmic Dynein at the Cytoplasmic Face of the ER Component of ES. Shown here are electron micrographs of late-stage spermatids that were processed for standard EM and embedded in Unicryl. After ultra sectioning these grids were incubated with an anti-cytoplasmic dynein (IC74) antibody followed by an anti-mouse gold (10 nm) conjugate. Panel A. Shown here is a longitudinal high magnification micrograph of a late-stage spermatid. The ER cistern is indicated and is seen adjacent to the actin layer of ES. Gold dots indicate the location of the IC74 component of cytoplasmic dynein. The gold is particularly concentrated to the cytoplasmic face of the ER membrane. Magnification: 56,000 X, bar = 0.25 um. Panel B. Shown here is a cross section high magnification micrograph through the head of a spermatid. The gold dots are located along the cytoplasmic border of the ER membrane. Magnification: 56,000 X, bar = 0.25 um. Panel C. Shown here is a high magnification micrograph through the head and acrosomal regions of a late-stage spermatid. Note that the majority of gold particles are concentrated to the area adjacent to the ER. Magnification: 80,000 X, bar = 0.5 um. Panel D. Shown here is a high magnification micrograph of a late-stage spermatid. Gold dots are located adjacent to the cytoplasmic face of the ER membrane. Magnification: 72,000 X, bar = 0.5 um. Panel E. Shown here is a longitudinal high magnification micrograph of a late-stage spermatid. The gold dots in this figure outline the cytoplasmic face of the ER. The actin layer is clearly visible between the ER and the plasma membrane of the Sertoli cell. Magnification: 28,000 X, bar = 0.5 um. 72 1 13 Figure 12. Pre-embedded Immuno-gold targeting of the IC74 Component of Cytoplasmic Dynein to the Manchette Region of the Spermatid. Panel A. Shown here is an EM of a cross section through the manchette region, of a spermatid embedded in unicryl. The manchette region, located on the anterior side of the spermatid head, shows a strong presence of IC74 immuno-gold labelling. Panel B. Shown here is a cross section through the tail region of a spermatid showing a strong presence of IC74 immuno-gold labelling at the flagellar and axonemal regions. Magnification A: X 24,000, B: X 76,000, bar = 0.25 um 74 1-5 Discussion Spermatogenesis is an event that occurs in the testis and involves two major cell populations, namely: Sertoli cells and spermatogenic cells. During spermatogenesis, round spermatids mature, ultimately, to spermatozoa, which are elongate in shape. Elongate spermatids are attached to Sertoli cells at sites where ES's form around developing spermatids. During the maturation process spermatids are transported from the apex of the Sertoli cell to its basal regions and are then returned to the apex. Elongate spermatids are then released to the lumen of the seminiferous epithelium during spermiogenesis. Spermatids, during the basally directed translocation event, become positioned deep within crypts of the Sertoli cell. In contrast, during the apically directed movement of the spermatids, before their release to the lumen of the seminiferous tubule, the crypts become shallow. The reason for the spermatid entrenchement is unknown; however, it has been proposed (Beach et al. 1999) that this entrenchment may provide a means of communication between adjacent spermatids and Sertoli cells. Do ESs (associated with spermatids) support microtubule movement? To answer this question a motility assay was developed (Beach and Vogl, 1999). In this assay the experimental approach was to place spermatids onto a glass slide fitted with a flow chamber (special coverslip) and then to add to this flow chamber fluorescently labelled microtubules. With this protocol it became possible to search for, using a Zeiss Axiophot microscope with fluorescence capabilities, the movement of microtubules along the spermatid 76 head. The spermatids collected for this study were isolated from the seminferous epithelium as follows. The epithelium was squeezed out of the seminferous tubule wall and then this tissue was aspirated though a pipette to physically fragment the tissue. The ectoplasmic specializations remain attached to the spermatid head and in most cases even the ER component of the ES remains remarkably intact. Since microtubules are known to be present and arranged in a parallel fashion around the elongated apical crypts of Sertoli cells it has been hypothesised that microtubule motors are involved in the translocation process. It is postulated that the motors are anchored to the ER component of ES and move along the ER-associated adjacent microtubules. This model proposes that the motor proteins provide the force that would then be transferred to the spermatid, causing it to translocate with its ES. The specific aim of the motility assay was to determine if isolated spermatids could support the movement of microtubules in vitro. The results indicated that isolated spermatids do support microtubule movement in the appropriate conditions (Beach & Vogl, 1999). This data strongly supports the motor based spermatid translocation process. In the Beach and Vogl (1999) study several important results were attained. Firstly, they observed that microtubules move along junction plaques that are associated with spermatids. This observation supported their initial working hypothesis: 'that spermatid translocation in the seminiferous epithelium is microtubule-based and involves motor proteins that are anchored to the junction plaque'. Secondly, their hypothesis is also supported by the 77 arrangement (parallel to long axis of cell and minus-end at cell apex) and association of the microtubules with the ES. Lastly, the fact that microtubules moved along the spermatid head only when the appropriate concentrations of nucleotide were present strongly supports their hypothesis since motors are ATPases. The Beach and Vogl study also discussed the following observation: Spermatids which had a partially detached or damaged ES, only supported microtubule movement in those areas of the spermatid head that had an undamaged ES. This finding is important since it indicates that the ES is absolutely required for microtubule movement on the spermatid head (Beach and Vogl 1999). 78 Figure 13. - Schematic Diagram of the Relationship Between the ES and the Associated Cytoplasmic Dynein Motor Proteins. This schematic diagram illustrates the proposed structural relationships between the cytoplasmic face of the ER, the dynein motor protein and the ES-associated microtubules. The left side of the figure is a cross section through the head of an elongate spermatid, as it would appear deep in a Sertoli cell crypt, and thus ready to undergo an apically directed translocation. The upper right-hand side of the figure is a more detailed view of the relationship between the ER component of the ES, the dynein motor protein and the ES associated microtubules. The lower right-hand side of the figure is a drawing of the dynein motor protein presented by Hirokawa (1998) which details the various components of the protein. Note that the IC74 (BAbCo and Pfister) and DHC1a (Asai) antibodies, that were used for the immuno-EM studies, are specific to the intermediate chain and the heavy chain of dynein, respectively, as illustrated on this figure. 6 79 Proposed Association of ER with Cytoplasmic Dynein and with Microtubules IC74 Immuno-Reactivity is Associated with the Cytoplasmic Face of the ER Component of ESs In this study we present morphological data that indicates that cytoplasmic dynein (IC74) is localized to the cytoplasmic face of the ER component of ES. Although previous studies (Miller et al. 1999) have shown, using immuno-fluorescence, that cytoplasmic dynein (IC74) is present in the area immediately surrounding elongate spermatid heads, their methodology was unable to specifically localise the dynein motor proteins to the ER. The data collected in this EM study are consistent with the motility hypothesis since this data supports the prediction that dynein motor proteins mediate the apical translocation of spermatids. The motility hypothesis provides a plausible mechanism for the spermatid translocation. This model exploits the well-documented (Hirokawa, 1998) interaction of enzymatic motors with microtubules. Our motility hypothesis suggests that enzymatic motors are anchored to the cytoplasmic face of the ER and move along the adjacent microtubules. Microtubules in Sertoli cells are oriented such that their plus-ends are located basally and their minus ends-are located apically (Redenbach et al. 1991). Since maturing spermatids translocate in both directions, that is, to the apical and basal regions of the Sertoli cell, we predict that both minus-end and plus-end directed microtubule motors mediate the translocation of spermatids. Based on the cytoarchitecture of Sertoli cells and the fact that cytoplasmic dynein is an ubiquitous and multifunctional motor protein we are 81 proposing that cytoplasmic dynein is responsible for the apically directed translocation of spermatids. Control of Spermatid Translocation In rat, step 16 (stage III) spermatids are located in apical regions of Sertoli cells. These spermatids are then translocated basally to the nuclear region of Sertoli cells (Figure 6, panels 1 to 2). Because the initial basally directed event proceeds towards the plus-ends of ES-associated microtubules it is likely that a kinesin or kinesin-like motor mediates this phase of the translocation event. Prior to the translocation event, ES junctional complexes form around apically located (Stage IX) early elongating spermatids. During this junction assembly process, it is possible that both kinesin and dynein are recruited to and are anchored to the cytoplasmic face of the ER. In a study by Muresan et al. (1996) they presented evidence that vesicles travelling along microtubule tracks in squid axons are often endowed with both dynein and kinesin motors. The question that immediately arises is: How can a vesicle move if oppositely directed motors are tugging on it ? The explanation that they present, is that both classes of motors (kinesin and dynein) are initially anchored to the lipid membrane of the vesicle and that once the vesicle reaches the distal end of the axon, kinesin detaches from the vesicle (possibly via a phosphorylation event) allowing dynein to mediate the return of the vesicle to the cell soma. How does the vesicle move to the plus-end of the microtubule in the first place, with dynein also tugging on it? In the same paper, they suggest that kinesin overrides dynein when both motors are anchored to the 82 same cargo. They support this conclusion with data indicating that kinesin is more processive than dynein; that is that kinesin can move for hundreds of steps (several micrometers) along a microtubule without dissociating from the microtubule (Romberg, et al.; Hancock et al.), that it moves in a "non-jittery" path (dynein movements are often jittery and dynein often dissociates temporarily from the microtubule along which it is travelling) and that kinesin has a higher duty ratio (it spends a greater fraction of its cycle time in a strongly bound, force-generating state) as compared with dynein. They also mention that even if there was only one kinesin, but, ten dyneins anchored to a single vesicle that the vesicle would move in the kinesin direction. Thus, kinesin must detach from the vesicle to allow dynein to move the vesicle back to the cell soma. They do not address the question of how kinesin returns to the cell soma once it becomes detached from the vesicle. This model suggests that if kinesin is attached to vesicles (regardless of the presence of dynein) that the vesicles will move to the plus-end of microtubules. Using the Muresan et al. (1996) model to explain the spermatid translocation event may be appropriate, because just as vesicles are transported bidirectionally along squid axon microtubule tracks, so are spermatids moved bidirectionally along microtubule tracks. Consider that both classes of motors are initially anchored (dynein may be anchored via dynactin) to the cytoplasmic face of the ER of apical spermatids (Figure 14). Kinesin overrides the activity of dynein (Figure 14, panel 1 to 2) and the spermatids are translocated to the base of the Sertoli cell (step 17 (stage V) of spermatogenesis). Spermatids are then translocated apically (Figure 14, 83 panel 2 to 3) for spermiation (Figure 14, panel 4; step 19 (stage VIII)). This apically directed translocation event, as evidenced by the immunolocalization data in this thesis, is most likely mediated by cytoplasmic dynein. I predict that cytoplasmic dynein motors at Stage V of spermatogenesis mediate the apically directed translocation event. This prediction is consistent with the apically directed translocation phase, if kinesin dissociates (Figure 14, panel 2) from the cytoplasmic face of the ER (possibly via a phosphorylation event) while the spermatid is positioned basally. The recruitment and anchoring of motors to the cytoplasmic face of the ER is likely part of the assembly (Figure 14, panel 4) of the ES junctional plaque. Based on this model, dynein motors would be anchored to the ER even during the kinesin mediated plus-end directed phase of the translocation event (Figure 14, panel 1). During the apically directed translocation phase, kinesin would dissociate from the cytoplasmic face of the ER in order for dynein to mediate movement. A major problem, in my opinion, with this model is that it does not address the question of how the kinesin motors are recycled back to the apical region of the Sertoli cell (Figure 14, panel 4- lower arrow indicating the return of the motors to the junctional plaque assembly process). In another study, Haimo (1995) presents an alternative possibility of bi-directional and motor-based vesicle movement control. She suggests that the control of bi-directional movements of a vesicle along microtubule tracks is mediated by the activation and deactivation of kinesin (Figure 15). She argues that for the kinesin directed movement that kinesin may be phosphorylated and 84 thus, activated. The activation of kinesin would allow it to override the potential activity of dynein. Once kinesin reaches the end of the microtubule, kinesin is dephosphorylated (deactivated) and thus, dynein moves the vesicle (with the attached dephosphorylated and deactivated kinesin) back to the cell soma. In the spermatid translocation model, a similar type of control may occur. It is possible that kinesin is activated on spermatids located at the apical regions of Sertoli cells (Figure 15, panel 1) and that once the spermatid reaches the basal region of the Sertoli cell (Figure 15, panel 2) then kinesin becomes deactivated and thus, dynein is able to mediate the return of the spermatids to the apical region of the Sertoli cell for spermiation (Figure 15, panel 4). This model requires, as does Muresan et al.'s, that kinesin is able to override dynein. Haimo's model may be more appropriate as a model for spermatid translocation since it accounts for the recycling of the kinesin motors back to the apical region of the Sertoli cell (Figure 15, panel 2 to 3). An important and attractive feature about Haimo's model is that the motors are consistently anchored to the ER. This is important because it may partly explain how the motors (both kinesin and dynein) associated with ES junctional plaques, that have disassembled, prior to spermiation become situated at the apical regions of the Sertoli cell. Thus, these apically situated motors may then be re-assembled into new ES plaques around other spermatids that have not yet undergone the translocation event (Figure 15, lower part of panel 4). 85 A third model to explain the bi-directional movements characteristic of the spermatid translocation event may simply be a stage specific recruitment of either kinesin or dynein motors to the ES junctional plaque (Figure 16). In a study conducted in our lab (Miller et al. 1999) immuno-fluorescence microscopic evidence was presented that indicated that apically positioned early elongate spermatids (confirmed by the presence of manchette) did not stain nearly as strongly for dynein (IC74 component) as compared with spermatids that were positioned basally (Stage V). This data suggests that dynein may not be anchored to the ES junctional plaques until the spermatid reaches the basal regions of the Sertoli cell. Thus, the data collected in this study forms the basis of the following model. Kinesin, may be recruited to and anchored to the cytoplasmic face of the ER of elongating spermatids that are positioned at the apical regions of Sertoli cells. Kinesin would then translocate spermatids along microtubule tracks and position them basally (Figure 16, panel 1 to 2). Dynein would then be recruited (mechanism unknown) to the junctional plaque and become anchored to the cytoplasmic face of the ER. As discussed above, it is likely that kinesin motors detach (Figure 16, panel 2) from the ER to allow dynein to mediate the minus-end directed translocation of the spermatid (Figure 16, panel 2 to 3). The spermatid would then be positioned apically in order for spermiation to occur (Figure 16, panel 4). The question that arises with this model is: How are the motors transported back to the regions of the Sertoli cell in which they anchor to the cytoplasmic face of the ER? It is possible that the motors are recycled and repositioned back to their 86 sites of attachment to the ES by being transported by oppositely directed motors or perhaps the motor is simply degraded once it reaches the end of the microtubule. This model is different from the above described models because in this case only one class of motor is anchored to the ES junctional plaque at any given phase of the translocation event. Thus, this model does not need to account for the added complexity of how oppositely directed motors are able to overcome each other during their respective active phases. CLIPS May Link Motors to ES CLIP-170 and other cytoskeletal linkers may play a role in the positioning and anchoring of the dynein motors for optimal contact with both the microtubules and the ER. Subsequent to the release of elongated spermatids to the lumen of the seminiferous tubule it is possible that dynein motors, as proposed above, along with linkers such as CLIPs, are recycled by being recruited to sites of ES assembly around apically located elongating spermatids. The isolation and collection of tissue to be studied using Immuno-EM required that the seminiferous epithelium be fragmented. The fact that the dynein motors remained anchored to the cytoplasmic face of the ER despite of the fragmentation procedure, suggests that the motors are firmly anchored to the cytoplasmic face of the ER and further supports this study for the following reason. The translocation of elongate spermatids, which are large cells, along microtubule tracts requires a considerable amount of locomotive force (piconewtons). Clearly, many motors as well as a very secure attachment of 87 the motor to the cytoplasmic face of the ER, would be required in order to transport the spermatid. The ER membrane, would of course, also need to be tear resistant, which seems to be the case since it remains intact after the fragmentation procedure. 88 Figure 14. - Schematic Diagram Illustrating a Mechanism of Control of the Bi-directional Control of Spermatid Translocation. A four panel schematic diagram (panel one at left) illustrating the phases of the spermatid translocation event (modified from Vogl et al. 2000). This particular model (the arrangement and organisation of the kinesin and dynein motors) is based on a model proposed by Muresan et al. (1996) to explain the translocation of vesicles along squid axons. In panel one both kinesin and dynein are anchored to the cytoplasmic face of the ER which is adjacent to the microtubules. Since kinesin "overrides" dynein, the spermatid is moved to the plus-end of the ES associated microtubules and becomes positioned basally (panel 2). Kinesin then detaches from the ES junctional plaque (mechanism unknown) to allow dynein to mediate the apical translocation of spermatids (panel 3). The apical junctional plaque then disassembles (panel 4) to allow for spermiation. As indicated by the arrows in panel 4 it is possible that the motors and the junctional plaque components are recycled (mechanism unknown) to regions of the Sertoli cell in which elongating spermatids are developing. 89 Figure 15. - Schematic Diagram Illustrating a Mechanism of Control of the Bi-directional Control of Spermatid Translocation. A four panel schematic diagram (panel one at left) illustrating the phases of the spermatid translocation event (modified from Vogl et al. 2000). This particular model (the arrangement and organisation of the kinesin and dynein motors) is based on a model proposed by Haimo (1995) to explain the bi-directional movement of vesicles. In panel one both kinesin and dynein are anchored to the cytoplasmic face of the ER which is adjacent to the microtubules. Since kinesin "overrides" dynein (when phosphorylated), the spermatid is moved to the plus-end of the ES associated microtubules and becomes positioned basally (panel 2). Kinesin then becomes dephosphorylated (deactivated-mechanism unknown), but remains attached to the ES junctional plaque (panel 2). Dynein then mediates the apical translocation of spermatids (panel 2 to 3). The apical junctional plaque then disassembles (panel 4) to allow for spermiation. As indicated by the arrow in panel 4 it is possible that the motors and the junctional plaque components are recycled (mechanism unknown) to regions of the Sertoli cell in which elongating spermatids are situated. 91 Figure 16. - Schematic Diagram Illustrating a Mechanism of Control of the Bi-directional Control of Spermatid Translocation. A four panel schematic diagram (panel one at left) illustrating the phases of the spermatid translocation event (modified from Vogl et al. 2000). This particular model (the arrangement and organisation of the kinesin and dynein motors) is based on data collected from a study conducted in our lab (Miller et al. 1999) suggesting that the dynein motors are more abundant in the basal regions of Sertoli cells. Thus, in panel one only kinesin would be anchored to the cytoplasmic face of the ER (adjacent to the microtubules). Kinesin then moves the spermatid to the plus-end of the ES associated microtubules (panel 2). Kinesin then detaches from the ES junctional plaque (mechanism unknown) while dynein is recruited to and anchored to the ES junctional plaque (panel 2). Dynein then mediates the apical translocation of spermatids (panel 2 to 3). The apical junctional plaque then disassembles (panel 4) to allow for spermiation. As indicated by the arrows in panel 4 it is possible that the motors and the junctional plaque components are recycled (mechanism unknown) to regions of the Sertoli cell in which elongating spermatids are situated. 93 Quantity & Density of Gold Particles Since the EM sections are only 70nm thick, each section only represents approximately .5% of the total length of a spermatid head. Thus if, for example, 10 gold particles are present on a micrograph (along the circumference of the cytoplasmic face of the ER), approximately 2000 gold particles would be expected to be present on the entire ER of the spermatid head (although, the IC74 complex is composed of three identical IC74 protein chains, it is unlikely, based on the size of the gold particle (10nm) that more than one gold particle would be able to bind to adjacent IC74 chains of a single motor). Thus, in order to gain an appreciation for the model that is being proposed it is essential that a three dimensional consideration of the system be appreciated. In a typical apically located spermatid perhaps 20-200 microtubules may be adjacent to the cytoplasmic face of the ER. Motors (as many as 2000) would then bridge the gap between the microtubules and the ER (cytoplasmic face) within the Sertoli cell. Motors would then need to move concurrently to transport the spermatid apically. CLIPs and Spermatid Translocation In our spermatid translocation model we propose that spermatids are translocated apically by cytoplasmic dynein along microtubule tracts. Consider, though, that initially this translocation machinery must be positioned with a specific orientation for an effective transportive event to occur. Thus, it is likely that CLIPs are involved in the spermatid translocation model since they may be required to ensure an optimal positioning between microtubules and cargo 95 (in this the ER). Thus, in this system CLIPs could be involved in correctly aligning the motors with the ER component of the ES and the adjacent microtubules. Possible Limitations of this EM study There are some limitations to the model proposed above. Firstly, a monoclonal antibody to the IC74 component, which is not the most highly conserved region of the molecule, was used to target cytoplasmic dynein. Notwithstanding this fact, I am not aware of any other family of proteins that contain an IC74 element. Even though the EM data indicates the presence of cytoplasmic dynein along the cytoplasmic face of the ER, this does not necessarily imply that cytoplasmic dynein is responsible for the apically directed spermatid translocation event. 96 CHAPTER 3 CYTOPLASMIC DYNEIN IS LOCALIZED TO THE ER COMPONENT OF ES: BIOCHEMICAL EVIDENCE Introduction In Chapter Two immuno-EM data was presented that indicated that cytoplasmic dynein is localized to the cytoplasmic face of the ER component of the ES junctional plaque. In this Chapter I present biochemical data that is consistent with our hypothesis and with the immuno-EM data. For these biochemical studies gelsolin treated tissue was used. Gelsolin is an actin-severing enzyme that dissociates, by means of actin digestion, the ER layer away from the ES. With centrifugation it is possible to concentrate the ER (supernatant) from spermatid heads and tails (pellets). The basic rationale for this experimental approach is to provide a method of comparing, using western analysis, the relative concentration of cytoplasmic dynein components in ER-enriched samples (supernatant) with ER-poor samples (supernatants of controls). This approach is appropriate since I am hypothesising that cytoplasmic dynein is highly concentrated at ER sites. The objectives for the biochemical study were as follows: 1) To confirm that the supernatants from gelsolin treated tissue are indeed enriched for ER. This was verified by probing supernatant-blots with an antibody specific to G R P 9 4 , which is, an ER resident protein. 2) To determine if ER enriched supernatants contain higher concentrations of IC74 (a component of dynein) as compared with supernatants that were not enriched for ER. 98 3) To determine if the ER enriched supernatants contain higher concentrations of DHC1a (a component of dynein) as compared with supernatants that were not enriched for ER. 99 Materials and Methods Testis Preparation for Gelsolin Experiment To obtain a sample of seminiferous epithelium enriched for elongate spermatids with attached junction plaques, the following protocol was followed: Testis were collected from rats in the same manner as described in the Material and Methods section of Chapter two. Testis were decapsulated, and the seminiferous tubules were placed into freshly prepared ice-cooled PEM/250 (80 mM PIPES, 1.0 mM EGTA, and 1.0 mM MgCI2, 250mM sucrose (pH 6.8 with KOH, also all working solutions contained 10 jag/ml soybean trypsin inhibitor, 0.5 |ug/ml leupeptin, 0.5 j^g/ml pepstatin, and 0.25 mM PMSF). Then using two sterile scalpel blades, the tubules were cut into small sections. The seminiferous epithelium was then squeezed out from the tubules with ultra-fine surgical tools (this was accomplished by arresting one end of the tubule with a fine-tipped probe and with a second probe, squeezing the epithelium out of the free end of the tubule) under a Zeiss dissecting microscope fitted with dark field optics. The collected epithelium was then transferred to a test tube containing ice-cooled PEM/250. This collection procedure continued for a maximum of 60 minutes. The collected tissue was then centrifuged at moderate speed in a clinical centrifuge for 2-3 minutes. The pellet was then re-suspended in 100-200ul of PEM/250 and gently aspirated (5-7 times) through a fine-bore pipette to mechanically dissociate spermatids from Sertoli cells. This tissue was then loaded to three step-sucrose gradient (60%, 55%, 50%, 45%, 40%, 35%, 30% sucrose in PEM) filled centrifuge tubes 100 (5 x41 mm-Beckman, Fullerton, CA) and centrifuged for 8.5 minutes at 5000 rpm in a Beckman SW Ti rotor. The band that formed in the Beckman tubes just below the 40-45% area, which was enriched for elongated spermatids with attached junctional plaques, was removed, using a syringe fitted with a 23G needle. Bands from three centrifuge tubes were pooled and placed into 1ml of ice-cold PEM/250. The tissue was further pelleted by centrifugation for 2-3 minutes at setting 6 on an Eppendorf table top (Hamburg, Germany) 5415C centrifuge, and then gently re-suspended in 50 ul of PEM/250. This method is similar to that detailed by Vogl (1996). Gelsolin Gelsolin is an 80 kDa protein which is a member of a class of actin modulating proteins. Gelsolin has powerful effects on actin filament length by exercising the following three modulating mechanisms: 1. Severing of actin filaments 2. Capping ends of actin filaments 3. Nucleation of actin filaments during assembly. It has been shown that gelsolin causes the formation of a large number of short actin filaments that are capped at their barbed ends (Kreis, 1993). The gelsolin family of proteins is one of the few known to sever actin filaments and thus, rapidly change actin filament length and number distributions. Ca 2 + promotes the interaction of gelsolin and actin whereas, polyphosphoinositides inhibit such interactions (Kreis, 1993). 101 Recently, several other actin severing proteins have been studied. Scinderin or adseverin is a 74-79 kDa Ca 2 + regulated actin filament severing protein found in adrenal chromaffin cells and other neuroendocrine tissues. It has a different amino acid composition than gelsolin, but, like gelsolin it is also inhibited by polyphosphoinositides. gCap39, is a 39 kDa protein that has extensive sequence homology to gelsolin, however, it only caps and does not sever actin filaments (Kreis, 1993). As explained further in this Chapter, gelsolin was used specifically as an actin severing enzyme. Since the ER component of ES is linked to the ES via actin filaments, an enrichment of the ER was obtained by treating the seminiferous epithelium with gelsolin and then using centrifugation to concentrate the ER fraction of ES from the rest of the tissue. Gelsolin has been used by others to sever actin filaments, for example, a study by Khaitlin and Hinssen (1997) showed that filamentous actin (ECP- (Mg) actin) was immediately disassembled by gelsolin when added at a 1:100 molar ratio. ER Enrichment Protocol Actin Severing Experiment (Gelsolin Experiment, Figure 17) Spermatids (with attached ESs) were obtained and suspended in MES (2-(4 morpholino)-ethane-sulfonic acid) buffer (50 mM MES-KOH pH 6.3, 2 mM MgCI2, 0.1 mM CaCI2 0.5 MM dithiothreitol [DTT]). The tissue was washed once in MES buffer, centrifuged at 6000 rpm for 2 minutes in an Eppendorf table top centrifuge, re-suspended in MES buffer and then divided into two equal aliquots (by volume). Both aliquots were then centrifuged at 6000 rpm for 102 2 minutes. One of the two samples (experimental) was re-suspended in gelsolin dialysate (0.4 mg/ml gelsolin in MES buffer) and the other (control) was re-suspended in MES buffer alone. Samples were frequently agitated and incubated for 60 minutes at room temperature. Small samples (5 ul) from each of the two incubations were collected and loaded onto polylysine-coated slides and stained with Oregon green-conjugated phalloidin (a fluorescence stain for actin filaments). These slides were then examined with fluorescence microscopy to verify that the gelsolin treated samples were, as compared to non-gelsolin treated samples, relatively void of actin filaments. Remaining samples were then centrifuged at 6000 rpm for 2 minutes to isolate the following four sample types: 1. Supernatant of gelsolin treated tissue 2. Pellet of gelsolin treated tissue 3. Supernatant of non-gelsolin treated tissue 4. Pellet of non-gelsolin treated tissue. Each of these samples were aliquoted (15 ul each) and stored at -70 °C. For this present study I did not probe samples two and four (pellets of control and experimental) with any of the antibodies. Gel Electrophoresis & Western Analysis SDS polyacrylamide gels followed by western blot analysis were performed to compare immunoreactivity of three proteins in supernatants of gelsolin and non-gelsolin treated fragmented seminiferous epithelium. The antibodies used were (1) mouse monoclonal anti-IC74 (BAbCo or Pfister) (2) 103 rabbit polyclonal anti-DHC1a (Dr. Asai) (3) rat monoclonal anti-GRP94 (Stressgen Biotechnologies). These antibodies were used to probe the above described tissue and compare immunoreactivity (gelsolin vs. non-gelsolin treated tissues) for cytoplasmic dynein (IC74 and DHC1a) and ER (GRP94). A Bio-Rad (Hercules, CA) Vertical Electrophoresis System (Protean II xi Cells- 16 x20 cm) with a 6.7% Laemmli resolving gel (Laemmli, 1970) was used to resolve tissue by molecular weight (for DHC1a and GRP94 experiments). Hoeffer Scientific Instruments (San Francisco, CA) mini gels (10 x 8.5 cm) were used for the IC74 experiment with 10.0% Laemmli resolving gels. The gels were prepared by mixing the components of the polymer and were then poured between two glass plates separated by 1 mm. A 1-2 cm layer of ddH 20 was applied to the top of gels to ensure that a straight interface between the resolving gel and the overlying stacking gel occurred. Experimental and control samples were diluted 1:1 with standard treatment buffer, boiled for 6 minutes at 110 °C (in dry bath) and then loaded to the gel wells at equal volumes and not at equal protein concentrations. The reason for loading equal volumes of samples, rather than equal protein concentrations, was because the supernatants of non-gelsolin treated samples were expected to contain less protein (since the entire spermatid was expected to be 'spun-down' to the pellet) as compared to the gelsolin treated samples. Volumes ranging from 10-40 pi were added to each well. The technical conditions that were most appropriate for this study, which required the resolution of high molecular weight proteins were as follows: 1 hour at 75 104 V/100mA / 1000 W, followed by 7 hours at 195 V /100mA /100W. The resolved proteins were then transferred to PVDF membranes using a Bio-Rad Trans-Blot Cell apparatus set to the following technical specifications: 4.5 hours at 10 V /100mA /100 W, followed by 1.25 hours a 75 V / 500 mA /100W. Membranes were then washed twice in TTBS (100 mM Tris-CI ph 7.5, 0.9% NaCI, 0.1% Tween 20) for 20 minutes each and then blocked (2 hours at room temperature) in 1:5000 normal goat serum (NGS), 4% bovine serum albumin (BSA), 1:5000 fetal bovine serum (FBS), 2% nonfat dry milk in TTBS. The series of washes was then repeated twice. Experimental blots were probed for 2 hours with freshly prepared DHC1a antibody (1:1,000 diluted in TTBS) or IC74 antibody (1:10,000 diluted in TTBS) or GRP94 antibody (1:5,000 diluted in TTBS). Control blots were probed (2 hours) under the same conditions with the equivalent protein concentration of normal rabbit serum (for DHC1a) or normal mouse IgG (for IC74) or normal rat IgG (for GRP94). Blots were then washed twice with TTBS and twice with TBS (20 minutes each wash). Experimental and control blots were then probed with secondary anti-rabbit horse-radish- peroxidase (HRP) antibody (for DHC1a) or anti-mouse HRP (for IC74) or anti-rat HRP (for GRP94) (Santa Cruz Biotechnology Inc., Santa Cruz, CA). Blots were then washed again, as above, and then treated with Amersham ECL solution (Amersham, Piscataway, NJ). Bio-Rad Laboratories Kaleidoscope Prestained Standards were used as molecular weight markers for all experiments. For the DHC1a experiment IgM was also used as crude high molecular weight marker (kDa approx. 1,200). Importantly, the IgM was 105 not incubated with the standard treatment buffer before the SDS-PAGE component of the experiment. Immunoblots were then wrapped in clear plastic and placed immediately in an autoradiography cassette to expose pre-cut sheets of Kodak X-OMAT film. Exposures ranging in time from 5 seconds to 10 minutes were performed. The film was processed with standard darkroom protocols. Immunoblots were scanned into digital format, cropped and re-sized using Adobe Photoshop 5 with care taken to ensure that the integrity of the data was not altered. Repetition of Western Analysis Experiments GRP94 This experiment was replicated twice using identical experimental conditions, as described in the preceding pages. Dynein- IC74 This experiment was replicated three times using identical experimental conditions in each case, as described in the preceding pages. This commercially available antibody has been appropriately tested and used extensively (standard immuno-fluorescence), in our lab and the results reported in this thesis were consistent with those reported by Miller et al. (1999). DHC1a Initially, I experienced some technical difficulties using this antibody. It is a polyclonal antibody and thus it reacted non-specifically at several molecular weights. However, after certain technical modifications over eleven trials, such 106 as, blocking the PVDF membrane in a cocktail composed of several blocking solutions, I was able to markedly reduce the non-specific background staining. Once the background staining was diminished I continued to replicate this experiment (five times), by trying varying concentrations and incubation times of the primary and the secondary antibodies. 107 Results Western blot analysis was used to determine if antibodies raised against IC74 and DHC1a (components of cytoplasmic dynein) and GRP94 (a luminal ER peptide) showed more immunoreactivity to gelsolin treated seminiferous epithelium as compared to non-gelsolin treated seminiferous epithelium. GRP 94 (Glucose Regulated Protein) Immunoblots presenting gelsolin treated tissue (supernatant) showed strong immunoreactivity (Figure 18, +gelsolin lane), at a molecular weight of 94 kDa, when incubated with the anti-GRP94 antibody. Immunoblots presenting non-gelsolin treated tissue showed extremely weak immunoreactivity (Figure 18, -gelsolin lane), at molecular weight of 94 kDa, when incubated with an anti-GRP94 antibody, indicating that the non-gelsolin treated tissue (supernatant) contained insignificant amounts of GRP94. Cytoplasmic Dynein- IC74 Immunoblots presenting gelsolin treated tissue (supernatant) showed strong immunoreactivity (Figure 18, + gelsolin lane), at a molecular weight of 74 kDa, when incubated with the anti-IC74 antibody. Immunoblots presenting non-gelsolin treated tissue (supernatant) showed weaker immunoreactivity (Figure 18, - gelsolin lane), at a molecular weight of 74 kDa, when incubated with the anti-IC74 antibody, indicating that the supernatant from the gelsolin treated sample is enriched for dynein. Cytoplasmic Dynein- DHC1a 108 Immunoblots presenting gelsolin treated tissue (supernatant) showed some reactivity (Figure 18, +gelsolin lane), when incubated with a polyclonal anti-DHC1a antibody, at a molecular weight of approximately 500-600 kDa. In addition, a band with molecular weight of approximately 180 kDa appeared on the experimental blots. This band also appeared on the non-gelsolin treated tissue (supernatant) blots (Figure 18, - gelsolin lane). There was considerable streaking and non-specific immunoreactivity on these blots which made them difficult to interpret. Control immunoblots (data not shown) showed little reactivity, when incubated with normal rabbit serum, at a molecular weight corresponding to the experimental immunoblots. 109 Figure 17. - Schematic Summary of the Gelsolin Experiment Schematic diagram summarising the gelsolin experimental methodology and the results obtained from this study. The left side of the figure summarizes the control conditions, whereas, the right side of the figure summarizes the experimental conditions. Notice that the only difference between the experimental and control conditions is the presence (experimental) or absence (control) of the gelsolin (actin severing) enzyme. Since gelsolin severs actin filaments, the hexagonally arrayed actin layer of the ES is digested which frees the ER component of the ES from the rest of the ES junctional complex. Western analysis was used to compare the presence of three proteins, namely: 1) GRP94 - an ER marker, 2) IC74 - a component of cytoplasmic dynein, 3) DHC1a - motor end of cytoplasmic dynein. 110 Gelsolin Experiment ER Membrane Centrifugation c o O Dynein Motors 03 -t—' CD E CD Q . X LU 94 k D a -74 k D a -200 k D a --GRP94 -IC74 •DHC1a Actin Layer Gelsolin Treatment Centrifugation Western Analysis 111 Figure 18. - Biochemical Data The Top Two Rows (labelled phase and actin): The two upper panels (labelled phase) are phase contrast images of typical spermatids (non-gelsolin treated = left, gelsolin treated = right). The second row of panels are fluorescence microscopy images of the same two spermatids stained with a fluorescent actin stain (Oregon green conjugated phalloidin). Note that the gelsolin treated spermatid (right) does not present any actin at the level of the ES since the actin layer was dissolved away by the gelsolin enzyme. The Lower Three Rows: The left most column of panels are the molecular weight markers (labelled mwt. markers) for the three experiments. The middle column are the western blots of non-gelsolin treated tissue, showing reactivity to the three antibodies, namely: GRP94, IC74 and DHC1a. The right most column are the western blots of gelsolin treated tissue, showing reactivity to the same three antibodies. Note that the GRP94 antibody and the IC74 antibody show a marked increase in reactivity in the gelsolin treated tissue as compared with the non-gelsolin treated tissue. 112 Discussion Cytoplasmic Dynein (IC74)- Immunoblots In this study I show, using biochemical techniques, that the IC74 component of cytoplasmic dynein is highly concentrated in samples of seminiferous epithelium that are enriched for ER (of ES). The EM studies, presented in Chapter Two, provide morphological evidence that dynein motors are localized to the cytoplasmic face of the ER. It was important to confirm, however, using another technique (biochemical technique), that dynein motors are highly concentrated specifically at the ER. The discrepancy between (Figure 18), the concentration of dynein motor proteins associated with ER enriched samples of tissue as compared with non-ER enriched samples of tissue, provides compelling support for the immuno-EM data. Cytoplasmic dynein (IC74), which is predicted to be responsible for the apical translocation of spermatids, is highly concentrated at the ER in the immuno-EM studies (cytoplasmic face of ER) as well as in this biochemical study. Also, the biochemical data confirming the presence of a moderate concentration of dynein in non-ER enriched samples (a light band on Figure 18, IC74 panel, middle column) is consistent with the immuno-EM data since there are some gold particles in areas that are not associated with the cytoplasmic face of the ER (Figure 10B). Thus, it is not surprising that the IC74 biochemical data showed some positive reactivity to samples of tissue that were not enriched for ER. The reason for this reactivity, unrelated to ER-114 anchored dynein, was simply because of the fact that cytoplasmic dynein is also known to be present in other areas of the Sertoli cell (Criswell et al., 1998). Possible Limitation of Biochemical Study Gelsolin treated fragmented seminiferous epithelium is, by nature, a very complex tissue sample and even though the results presented in this thesis indicate that cytoplasmic dynein is highly concentrated at ER sites, it is possible that the positive immunoblot reactions are due to the abundant cytoplasmic dynein motors that are located elsewhere in the Sertoli cell or germ cell. Thus, the cytoplasmic dynein proposed to be responsible for the apically directed spermatid translocation process, may be cytoplasmic dynein that is required for instance, for transporting vesicles along the ES-associated microtubule tracts and thus, not related to spermatid translocation. From this possible limitation I conclude that it would be important to design an experiment to determine if the cytoplasmic dynein motors that have been shown to be associated with the ER, are indeed responsible for spermatid translocation. The following experiment, which may not be technically possible yet, may be able to determine if cytoplasmic dynein causes the apically directed translocation of spermatids. Isolated spermatids, labeled with a fluorescent ER stain (such as DiOC6), could be added to a slide "carpeted" with polarity labeled microtubules. Then if the spermatids were observed to move along these microtubules, in dynein direction, then a stronger functional association of cytoplasmic dynein with the spermatid translocation process could be proposed. This study would confirm that spermatids (with an attached ER) 115 move along microtubules, which is what is proposed to happen in vivo, rather than showing, as did the Beach et al., (1999) study, that microtubules move along spermatid heads. Other possible experiments to test the proposed requirement of cytoplasmic dynein in the spermatid translocation process involve the use of 'modified' motility assays. If binding of ER-anchored dynein motors to microtubules was inhibited, by blocking the active site on the dynein heavy chains with a specific antagonist, one would expect to observe a decrease in the movement of labelled polar microtubules (compared with non-blocked dynein motors), in the appropriate direction, along spermatid heads. This approach may be useful in confirming that labelled polar microtubules added to isolated spermatid junctional plaques are translocated specifically by cytoplasmic dynein motors. Another possibility to confirm that dynein is responsible for the translocation event would be to modify the 'motility assay' whereby spermatids that translocated microtubules, in the dynein direction, along the ES junctional plaque, were immediately processed for immuno-EM. In this approach these particular spermatids would be expected to present a complete ES junctional complex (especially the ER) and a strong positive stain (gold particles), along the cytoplasmic face of the ER, when probed with the anti-IC74 antibody and appropriate secondary antibodies. 116 Cytoplasmic Dynein - DHC1a In this study we show that DHC1a is concentrated in samples of seminiferous epithelium that are enriched for ER. Previous studies have shown that DHC1a is present in rat testis (Criswell and Asai, 1996). This finding coupled with the IC74 EM data strongly suggests that cytoplasmic dynein is localized to cytoplasmic face of the ER component of ES. Since the DHC1a antibody reacted with multiple bands on immunoblots, immuno-EM would not be expected to provide results comparable to the IC74 study and thus, EM experiments were not performed with the DHC1a antibody. This data is consistent with the other studies presented in this thesis and supports the hypothesis that cytoplasmic dynein is responsible for the apical translocation of spermatids. DHC1a is the 'engine' of cytoplasmic dynein and would be required for translocation to occur along microtubules and thus, is likely to be a component of the motor responsible for apically directed spermatid translocation. Although there are other isoforms of the dynein heavy chain in rat testis, DHC1a is the only one known to associate with the IC74 intermediate chain of cytoplasmic dynein. A recent study (Basak et al., 1999) described a technique that may be used in conjunction with western analysis to confirm the specificity of results obtained with my study. In their study, they incubated PVDF membranes (containing resolved proteins) in a medium that contained a molecule that blocked the antigenic site of the protein of interest. They subsequently incubated the immunoblot with the primary antibody (that was used for the 117 initial western analysis). The primary antibody, because the antigenic site was blocked, was unable to bind to the protein of interest and, a negative result was obtained. In my study a similar experiment could be performed to determine if the band on the DHC1a blot was indeed specific. The antigenic site of DHC1a would need to be blocked on the blot and then the blot would be probed with the DHC1a antibody. In this scenario a negative result would be expected if the preliminary results, presented in this thesis, are specific. A study such as this would be very useful to conclusively determine if DHC1a is indeed concentrated to the ER of the ES. Immunoprecipitation could also be performed to determine if the IC74 and the DHC1a components are co-localized at the level of the ER. GRP 94 The results of this study indicate that GRP 94 (glucose regulated protein) is highly concentrated in supernatants from gelsolin treated spermatid/junction complexes treated with gelsolin. GRP 94 is a soluble protein that is present in ER cisterns. GRP 94 is a member of the 90-kDa molecular chaperone family of proteins which comprises, among other proteins, the 90-kDa heat-shock protein, hsp90 and the 94-kDa glucose- regulated protein, major molecular chaperones of the cytosol and proteins of the endoplasmic reticulum (Csermely et al., 1998). My data are consistent with the conclusion that gelsolin released the ER from ES attached to spermatid heads. 118 The data presented in this thesis are consistent with previous studies which indicated the presence of cytoplasmic dynein at sites associated with spermatid heads located in Sertoli cell crypts (Miller et al., 1999). In addition, the data is supportive of the spermatid translocation model which proposes a major role for cytoplasmic dynein with respect to the apically directed translocation of spermatids that occurs during spermatogenesis. Overall Conclusions: 1) The immuno-EM data is consistent with the conclusion that cytoplasmic dynein is present on the cytoplasmic face of the ER component of ESs. In addition, cytoplasmic dynein is also found generally in Sertoli cell cytoplasm surrounding apical crypts and in stalks supporting late spermatids. Its presence in these locations may be related to the transport of membraneous organelles other than the ER of ESs. 2) The biochemical data was supportive of the conclusion indicated above and provided further evidence indicating a strong presence of cytoplasmic dynein (especially the IC74 study) at the level of the ER. 3) Based on the data presented in this thesis, cytoplasmic dynein may be responsible for the apically directed translocation of spermatids observed during spermatogenesis. However, the results do not preclude the possibility that a minus-end directed kinesin may be associated with the ER. The data also do not demonstrate that dynein actually moves the plaque in vivo. 119 Bibliography: Ahmad, F.J., et al. (1998): Cytoplasmic Dynein and Dynactin Are Required for Transport of Microtubules into the Axon, The Journal of Cell Biology, vol.: 140-2, :391-401. 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