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Plus-end and minus-end directed motors are associated with specialized Sertoli cell junction plaques… Guttman, Julian 1999

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f Plus-end and Minus-end Directed Motors are Associated with Specialized Sertoli Cell Junction Plaques (Ectoplasmic Specializations): Evidence From an In Vitro Polarity Marked Microtubule Motility Assay by Julian Guttman B.Sc., The University of Western Ontario, 1997 A THESIS S U B M I T T E D IN P A R T I A L F U L F I L M E N T O F T H E R E Q U I R E M E N T S F O R T H E D E G R E E OF Master of Science in T H E F A C U L T Y OF G R A D U A T E STUDIES (Department of Anatomy) We accept this thesis as conforming To the required standard, T H E U N I V E R S I T Y OF BRITISH C O L U M B I A September 1999 © Julian Andrew Guttman, 1999 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. Department of /jn*Aon\<j The University of British Columbia Vancouver, Canada Date Sspi. /£f£f*j DE-6 (2/88) Abstract Developing sperm cells (spermatids) translocate within the seminiferous epithelium during spermatogenesis. It has been proposed that this translocation results from the microtubule-based transport of specialized junction plaques that occur in Sertoli cells in regions where these cells are attached to spermatids. These junction plaques each consist of a cistern of endoplasmic reticulum, a layer of actin filaments, and a region of the plasma membrane involved with intercellular adhesion. Molecular motors anchored to the cytoplasmic face of the endoplasmic reticulum are thought to move the junction plaques, and hence the attached spermatids, towards the base and back to the apex of the epithelium, along microtubules. In Sertoli cells, microtubules are arranged parallel to the long axis of the cell and have their minus ends located at the apex of the epithelium. If the spermatid translocation hypothesis is true, then isolated junction plaques should support microtubule movement both in the plus and minus direction when tested in vitro. To verify this prediction, I have developed and have used an in vitro polarity-marked microtubule motility assay. Polarity labeled microtubules were bound to isolated spermatid/junction complexes and then the complexes were added to motility chambers. The complexes then were assessed for their ability to transport microtubules in the presence of a motility buffer containing ATP and testicular cytosol (2 mg/ml). The presence of junction plaques attached to spermatids was established with fluorescently labeled phallotoxin for filamentous actin, and with DIOC6 for endoplasmic reticulum. Of 25 recorded motility events, 17 were in the plus-end direction and 8 were in the minus-end direction (on a bed of purified kinesin 35/37 polar microtubules moved in the plus-end direction and on a bed of testicular cytosol all 158 polar microtubules moved in the ii plus-end direction). The results are consistent with the conclusion that the junction plaques have the potential for moving along microtubules in both the plus and minus directions. Also, together with some of my data from immunofluoresence studies, they constitute the first evidence that a kinesin-type motor is associated with the junction plaques at stages when translocation occurs in vivo. Table of contents Abstract ii List of Figures viii List of Abbreviations x Acknowledgements xi Introduction 1 The Organization of Spermatogenic Cells in Epithelia and their Relationship with Sertoli Cells 2 Microtubule-based Spermatid Translocation 4 Background 6 Ectoplasmic specializations 6 Microtubules in general 8 Dynamic instability model 9 Microtubules in Sertoli cells 14 Precedent for microtubule-based transport and pattern of the Endoplasmic reticulum in General 15 Conclusion 17 Endoplasmic reticulum imaging 18 Motors general 20 \ Kinesin 21 Dynein 24 Multiple motors 27 Motors in Testis 28 Kinesin in Testis 28 Kinesin in Sertoli Cells 30 Dynein in testes 31 Dynein in Sertoli Cells 33 Dynein in Spermatogenic Cells. 34 The microtubule-based spermatid translocation hypothesis 35 Thesis Problem (Hypothesis) 37 iv Materials and Methods 50 Animals 50 Chemicals and Reagents 50 Polar Microtubule Construction 50 Cytoskeleton Method of Polar Microtubule Construction 51 N E M method of Polar Microtubule Construction 53 G M P C P P Method of Polar Microtubule Construction 58 Preparation of Testicular Cytosol 61 Spermatid and Ectoplasmic specialization isolation 63 Preparing the perfusion chamber 65 The motility assay 66 Verification that ectoplasmic specializations were attached to spermatids... 67 Controls 68 Data Analysis 75 Rates 75 Statistics 75 Immunofluorescent Experiments for Kinesin 76 Preparation of Tissue Sections 76 Fixation for sectioned material 76 Sectioning 76 Preparation of isolated ectoplasmic specializations from tissue for Kinesin immunofluorescence 77 Immunofluorescence of Kinesin 78 Immunofluorescent Controls for Kinesin 79 Results 8 0 Polar Microtubules 80 Polar Microtubules 80 Cytoskeleton Kit 80 N E M Polar microtubules 81 Polar microtubules are in the appropriate direction; cytosol increases rates 82 G M P C P P Polar Microtubules 82 Cytosol supports movement in the plus-end direction 84 Polar Microtubules Move in both the plus and minus-directions on Isolated Ectoplasmic Specializations 86 Movements Trailing by the Seed 87 Movements Leading by the Seed 87 Kinesin Immunofluorescence 88 Controls 88 Discussion 119 Establishment of the assay to verify the prediction 120 Polarity-marked microtubules 122 Ectoplasmic specializations attached to isolated spermatids translocate Polar microtubules both in the plus and minus directions 124 Temperature 126 Evidence that the observed microtubule transport occurs on ectoplasmic specializations and is specific 127 Evidence from immunofluorescence indicates the presence of a kinesin-like motor at ectoplasmic specializations 129 Summary 130 Conclusions 131 Future Directions 132 References 133 vi List of figures Figure 1 Diagram of the seminiferous epithelium 38 Figure 2 Single panel diagram of the microtubule-based spermatid translocation hypothesis 40 Figure 3 Experimental model 42 Figure 4 Diagrammatic representation of microtubule polarity in cells 44 Figure 5 Four panel diagram of the microtubule-based spermatid translocation hypothesis 46 Figure 6 Direction of movement of anchored and un-anchored molecular motors and polar microtubules 48 Figure 7 Polarity marked G M P C P P microtubules bound to the ectoplasmic specialization 71 Figure 8 Experimental method 73 Figure 9 Microtubules constructed using the "Cytoskeleton" method 89 Figure 10 N E M non-polarity marked microtubules 91 Figure 11 N E M polarity marked microtubule seeds break apart 93 Figure 12 N E M polarity marked microtubule seed sticks to glass 95 Figure 13 N E M polarity marked microtubules capable of motility 97 Figure 14 Collage of G M P C P P polarity marked microtubules 99 Figure 15 G M P C P P polarity marked microtubule kinesin controls 101 Figure 16 G M P C P P polarity marked microtubule testicular cytosol controls. 103 Figure 17 Graph of polar microtubule motility rates 105 vii Figure 18 Plus-end directed movement on an ectoplasmic specialization . . . . 107 Figure 19 Minus-end directed movement on an ectoplasmic specialization... 109 Figure 20 Kinesin immunofluorescent staining on testis sections I l l Figure 21 Kinesin immunofluorescence on isolated spermatids with attached ectoplasmic specializations 113 Figure 22 Controls of Kinesin immunofluorescent sections 115 Figure 23 Controls of Kinesin immunofluorescence on isolated spermatid with attached ectoplasmic specializations 117 viii List of Abbreviations D M S O Dimethyl Sulfoxide PIPES Piperazine-N,N'-bis[2-ethanesulfonic acid] E G T A Ethylene Glycol bis[succininidyl succinate] MgC12 Magnesium Chloride Taxol Pacitaxel from Taxus yannanensis N E M N-ethylmaleimide G T P Guanosine 5'-Triphosphate E G S Ethylene glycol-bis-succinimidylsuccinate G M P C P P guanylyl (alpha,beta)methylenediphosphonate P M S F Phenylmethylsulfonylfluoride D T T Dithiothreitol D iOC6 3,3'-dihexylocarbocyanine iodide A T P Adenisine 5'-Triphosphate Tween-20 Polyoxylethylenesorbitan Monolautate FITC Fluorescein isothiocyanate NaCl Sodium Chloride KC1 Potassium Chloride Na2HP04 Disodium Phosphate K H 2 P 0 4 Monopotassium Phosphate PBS Phosphate Buffered Saline T P B S / B S A Tween/Phosphate Buffered Saline/. 1% Bovine Serum Albumin ix Acknowledgements I would like to thank Dr. Johnathan Howard for the gift of G M P C P P and his insight into N E M tubulin and polar microtubule polymerization, Kara Gabriel for her help with the statistical tests, Craig Martin for all of his computer assistance, Tara Callan for her help with various computer programs and Dr. Donald Moerman and Dr. Calvin Roskelley for their scientific guidance. I would also like to extend a special thanks to Dr. A . Wayne Vogl for his undying enthusiasm with regard to my project and for all of his support throughout all of my thesis work. Introduction In this thesis I investigate one aspect of the mechanism by which developing sperm cells (spermatids) are moved between the apex and the base of the seminiferous epithelium during spermatogenesis in mammals. The mechanism involves the translocation of specialized junction plaques (ectoplasmic specializations) by molecular motors along microtubules in Sertoli cells. Generally in these cells, as in most cells, cytoplasmic microtubule-based transport is associated with the intracellular movement of membranous organelles (Johnson et al., 1996). In order to use this system to move an adjacent cell, the Sertoli cell has incorporated a cistern of endoplasmic reticulum into the submembrane junction plaque of the Sertoli cell in regions of adhesion to spermatids. In this thesis I test the hypothesis that these junction plaques have the potential for moving along microtubules both in the plus and in the minus direction, as would be predicted by the microtubule-based spermatid translocation model. This is the first system in which movement of one cell by another cell has been shown to be accomplished through the use of microtubule-based molecular motors. l The Organization of Spermatogenic Cells in Epithelia and their Relationship with Sertoli Cells Sertoli cells and the developing spermatogenic cells together make up the seminiferous epithelium. Sertoli cells are columnar cells, which span the distance from the outer border of the seminiferous tubule (basal lamina) to the lumen of the seminiferous tubule and in doing so, support the developing spermatogenic cells (Figure 1). Within the cytoplasm of these cells, as with virtually all animal and plant cells, is a cytoskeletal scaffolding composed of actin filaments, microtubules, and intermediate filaments (Fuchs and Cleveland, 1998). The cytoskeletal system in general consists of the "muscles and bones" of a cell and is responsible for giving cells their shape, for moving organelles from one region of the cell to another, and for allowing cells to move within and between tissues. In order for the spermatogonia to become spermatozoa, they must undergo the complex process of spermatogenesis. Spermatogenesis involves the division and differentiation of spermatogenic cells into spermatozoa (sperm). Male stem cells, termed spermatogonia, proliferate through multiple divisions. Once they become spermatids they differentiate into sperm through a process termed spermiogenesis. From beginning to end spermatogenesis takes 57 days in rats and 64 days in humans (LeBlond and Clermont 1952). 2 Located along the outer borders of the Sertoli cells towards the base of the seminiferous epithelium are the most juvenile spermatogenic cells, spermatogonia. These spermatogonia are capable of dividing and producing daughter cells, some of which undergo further differentiation into spermatocytes (LeBlonde and Clermont, 1952). The spermatocytes undergo meiosis and thereby reduce the number of chromosomes and D N A in the cells. These cells dissociate from the basal lamina and move through basal junctions between Sertoli cells and enter the adluminal compartment of the epithelium. After passing into the adluminal compartment, the spermatocytes complete meiosis and give rise to diploid spermatids. These spermatids are initially round cells. They gradually polarize, undergo a complex process of morphological differentiation (spermiogenesis) and are ultimately moved to the apex of the Sertoli cells where they are released as fully developed sperm cells. 3 Microtubule-Based Spermatid Translocation One of the most dramatic events that occurs during spermatogenesis is the movement of the developing spermatids between apical and basal regions of the seminiferous epithelium (spermiogenesis) (Figure 2). At the apex of the epithelium, the spermatids become contained within depressions in the apical surfaces of Sertoli cells termed the apical crypts. These crypts deepen at one point during spermiogenesis and then again become shallow just prior to the spermatid release into the lumen of the seminiferous tubule. When this occurs, spermatids are moved from the apex of the Sertoli cell towards the base of the epithelium within apical invaginations (or crypts) formed parallel to the longitudinal axis of the Sertoli cell (Vogl et al., 1991). Following this, the spermatids are then returned to the apex of the epithelium where they are eventually released into the lumen of the seminiferous tubule (Vogl et al., 1991). Specialized Sertoli cell junction plaques termed ectoplasmic specializations occur in regions of the crypt adherent to the spermatid heads. These plaques are composed of the Sertoli cell plasma membrane, a layer of hexagonally packed actin bundles and a cisternae of endoplasmic reticulum, and are thought to be involved in the process by which spermatids are translocated in the epithelium (Grove and Vogl , 1989; Vogl, 1996). It has been proposed that these plaques have molecular motors attached to the cytoplasmic side of the endoplasmic reticulum component of the plaque and use the Sertoli cell's microtubules as tracts to translocate the developing spermatids both towards the base and the apex of the Sertoli cell (Redenbach and Vogl .,1991; Vogl et al., 1991). 4 In the following sections I will present background information related to ectoplasmic specializations, microtubules, associations of the endoplasmic reticulum with microtubules, and molecular motors. 5 Background Ectoplasmic specializations Specialized junction plaques, termed "ectoplasmic specializations", occur in regions where the Sertoli cells are attached to developing sperm cells (spermatids). These ectoplasmic specializations are situated adjacent to spermatids and are related both to Sertoli cell microtubules and to intermediate filaments during spermiogenesis (Vogl, 1989). Ectoplasmic specializations are composed of a layer of actin filaments which is linked to a cisternae of smooth endoplasmic reticulum on one side, and the Sertoli cell plasma membrane on the other (Vogl et al., 1991). This actin layer is about four to six filaments thick and the filaments are hexagonally packed. Molecular components identified at the site include actin (Toyama, 1976; Franke et al., 1978; Vogl et al., 1983; Suarez-Quian and Dym, 1984; Vogl and Soucy, 1985a; Vogl and Soucy, 1985b Vogl et al., 1986; Camatini et al., 1986; Camatini et al., 1987; Masri et al., 1987; Suarez-Quian and Dym, 1988; Fouquet et al., 1989;), a-actinin (Franke et a l , 1978; Jockusch and Isenberg, 1981), vinculin (Grove and Vogl , 1989), espin (Bartels et al., 1996), fimbrin (Grove and Vogl , 1989), myosin V i l a (Hasson et al., 1995) and a6(31 integrin (Pfeiffer and Vogl. , 1991). Towards the cytoplasm of the Sertoli cell lies the endoplasmic reticulum component of the plaque, the internal space of which actually appears thinner than any other endoplasmic reticulum found within the cell (unpublished observation). This organelle is linked to the actin layer, which in turn is attached to the plasma membrane overlying the acrosome of the spermatids (Figure 3). Because of the observed cross-links between 6 ectoplasmic specializations, specifically the endoplasmic reticulum component of the plaque, and neighboring microtubules, ectoplasmic specializations have been proposed, for many years, to be involved in positioning spermatids within the seminiferous epithelium (Christensen 1965; Fawcett 1975; Russell 1977; Vogl , 1988; Redenbach and Vogl 1991). When spermatids are mechanically displaced from the seminiferous epithelium, ectoplasmic specializations remain attached to the spermatid heads (Vogl, 1996). Significantly, the plaques remain intact; that is the plasma membrane of the Sertoli cell, the layer of actin filaments, and the endoplasmic reticulum can all be identified as present. These observations indicate not only that the major components of the plaque are a structural and probably functional unit, but also establish the basis for using mechanically dissociated spermatid preparations as an in vitro model for studying ectoplasmic specializations. 7 Microtubules in General Microtubules are one of the major classes of cytoskeletal elements found in cells. Their walls are usually composed of 13 parallel 4-5nm pro to filaments (Andre and Thiery, 1963; Pease, 1963; Fujiwara and Tilney, 1975), which in turn are formed of 8nm (in length) heterodimers. Each heterodimer is composed of an a and P tubulin protein each of 50,000D (Fujiwara and Tilney, 1975). The protofilaments together form a circular structure, giving the microtubule an overall diameter of about 24nm. The assembly of microtubules is G T P dependent (Olmsted and Borisy, 1975). G T P binds both to the a and to the p subunits of the heterodimer. During microtubule assembly, the G T P found on the p tubulin subunit is hydrolyzed to G D P plus phosphate after binding to an adjacent dimer has occurred. This makes the polymer less stable. The use of a non-hydrolizabe analogue of G T P (for example G M P C P P ) does not allow hydrolysis to occur; in other words, heterodimer units are inhibited from depolymerizing once they have polymerized to form a protofilament. When microtubules polymerize they do so in both directions. Instead of starting from one heterodimer and growing in one direction, microtubules begin their growth from heterodimers that have already bound together and grow outwards in both directions leaving the initial tubulin "seed" within the polymerized microtubule. When this elongation process occurs, the tubulin heterodimer subunits polymerize onto the nucleating centers 2-4 times faster on one end than the other. The side that is polymerizing faster is called the (plus) or fast growing end of the microtubule and the 8 side of the microtubule growing at a slower rate is called the (minus) or slow growing end. Microtubule polarity in cells determines the types of motors associated with transporting cargo to specific sites in the cell. The difference in polymerization kinetics is also essential to the construction of polarity marked microtubules used in experimentation, which will become apparent later in this thesis. Dynamic instability model Dynamic instability describes the behavior of microtubules where individual microtubules (independently of their neighbors) exist in phases of polymerization and depolymerization by the addition or loss of subunits from the microtubule ends (the more active end is termed the plus (+) end and the less active end is termed the minus (-) end) (MacRae, 1992; Cassimeris, 1993). These episodes of polymerization and depolymerization occur with infrequency (Cassimeris, 1993). Two main phases occur, the transition from elongation to rapid shortening which has been termed catastrophy, and the transition from rapid shortening to elongation, termed rescue (Walker et al., 1988; MacRae, 1992). Although not completely understood, dynamic instability is thought to be an intrinsic property of the tubulin molecule itself. It is likely that microtubules are inherently unstable and that they are stabilized by a cap of stable subunits (Mitchison and Kirshner 1984). In this general type of model, tubulin subunits add to the elongating end of a microtubule while in the stable conformation, and these subunits then undergo a destabilizing conformational change some time after assembly (Cassimeris, 1993). 9 Catastrophy occurs by the stochiastic loss of the stable cap (either by dissociation or conformational change), while rescue occurs if the stable cap is able to re-form (Walker et al., 1988; Walker et al., 1991). Assembly of tubulin subunits is promoted by the binding of GTP to an exchangeable site (E-site) on the tubulin dimer (tubulin itself is a GTPase) (Cassimeris et al., 1987; Erickson and O'Brien, 1992). That is, tubulin with its associated molecules of G T P is inserted into the growing end of the elongating microtubule (MacRae, 1992). Some time after this incorporation the G T P is hydrolyzed to G D P (MacRae, 1992) resulting in a "core" of tubulin-GDP and a "cap" of tubulin-GTP at the elongating microtubule end (Cassimeris et al., 1987). Conversion from one polymerization state to another may therefore be regulated by the amount of tubulin-GTP at the plus end of the microtubule (MacRae, 1992). In conditions where polymerization is occurring, it is theorized that microtubules have a G T P cap at the plus end and as long as it is present, the microtubules are stable (MacRae, 1992). This is due to a principal put forward by Mitchison and Kirshner that the dissociation rate of tubulin-GDP is much faster than the rate for tubulin-GTP (Mitchison and Kirshner, 1984; Mitchison and Kirshner, 1986). Thus, during elongation, a "cap" of tubulin-GTP stabilizes an unstable tubulin-GDP "core" (Cassimeris, 1993). When hydrolysis of G T P catches up with elongation, the tubulin-GDP core is exposed and catastrophe results from the rapid rate of tubulin-GDP dissociation and the low rate of association of tubulin-GTP or tubulin-GDP (Cassimeris et al., 1987). The end result is a 10 rapid and extensive microtubule depolymerization (Cassimeris et al., 1987). Rescue occurs when a tubulin G T P is added to a rapidly shortening end, thus re-capping the unstable core (Cassimeris et al., 1987). A non-hydrolysable analogue of G T P , G M P C P P , apparently binds strongly enough to displace the nucleotide binding site (termed the E-site) G D P , and supports multiple cycles of assembly (O'Brien and Erickson, 1989; Sandoval and Weber, 1980). G M P C P P is incorporated stoicheometrically into microtubules and, because it is not hydrolyzed (Erickson and O'Brien, 1992 ), G M P C P P microtubules appear to be much more stable than those polymerized with G T P itself (Walker and O'Brien, 1988). This high degree of stability has proven useful in polymerizing the seeds of polar microtubules. The use of G M P C P P allows the seeds to remain stable while the dimly labeled plus and minus ends of the microtubule are polymerized onto it. Dynamic instability is regulated by a number of cellular factors, including the structural microtubule-associated proteins, which in turn are regulated by phosphorylation (Drewes et al., 1998). Microtubules interact with several non-tubulin proteins termed microtubule-associated proteins (MAPs) (MacRae, 1992). These microtubule-associated proteins can be divided into two groups - the structural and the dynamic microtubule-associated proteins (MacRae, 1992). The structural microtubule-associated proteins stimulate tubulin polymerization, enhance microtubule stability, and influence the spatial distribution of microtubules within cells (MacRae, 1992; Dreshel et al., 1992; Triczek et al., 1995; Kaech et al., 1996). The dynamic microtubule-associated proteins are motor 11 proteins and exploit existing microtubule polarity and patterns of organization to carry out directed translocation of subcellular components (MacRae, 1992). Structural microtubule-associated proteins are filamentous proteins that lack enzymatic (nucleotide hydrolyzing) activity and bind to microtubules reversibly with affinities in the low-micromolar range (Drewes et al., 1998). The best characterized microtubule associated proteins (MAP2 and tau) were initially isolated from brain tissue (Cassimeris, 1993). Both these microtubule associated proteins increase the rate of microtubule elongation and increase the rescue frequency, but they also suppress the catastrophy frequency and reduce the shortening rate (Dreshel et al., 1992; Preyer et al., 1992; Kowalski and Williams, 1993). Thus, these types of microtubule associated proteins would not generate the dynamic microtubules observed in proliferating cells (Cassimeris, 1993). Members of the microtubule-associated protein family also include M A P 4 , which is present in all non-neuronal vertebrate cells (Mandelkow and Mandelkow, 1995; Schoenfeld and Obar 1994). A l l three of the previously named proteins are composed of an N-terminal projection domain, which protrudes from the microtubule surface, and a C-terminal microtubule-binding domain that contains a proline-rich sequence and three or four 31-residue pseudorepeats (Drewes et al., 1998). Overexpression of microtubule-associated proteins in cells can induce bundling of microtubules. The spacing of the microtubules depends on the projection domain of the transfected microtubule-associated protein (Chen et al., 1992). 12 The phosphorylation of microtubule-associated proteins is a crucial factor in the regulation of microtubule dynamics (Drewes et al., 1998). Phosphorylation of microtubule-associated proteins influences their microtubule-stabilizing capacity (Dreshel et al., 1992; Triczek et al., 1995; Biernat et al., 1993) and, accordingly, microtubule-associated proteins isolated from tissue (Matsuo et al., 1994) or cells (Illenberger et al., 1998) show varying degrees of phosphorylation (Drewes et al., 1998). Microtubule-affinity-regulating kinases were identified by their ability to destabilize microtubules by phosphorylating microtubule-associated proteins (Drewes et al., 1998). Microtubule-affinity-regulating kinases are novel mammalian serine/threonine kinases that phosphorylate the tubulin-binding domain of microtubule-associated proteins and thereby cause their detachment from microtubules and increased microtubule dynamics (Drewes et al., 1998). Summary Although dynamic instability is a concern when polymerizing microtubules, the use of non-hydrolyzable analogues of G T P (eg. GMPCPP) abolish this dynamic instability. In addition to this, stability to microtubules can be increased and dynamic instability can be controlled through post-translational modifications in vivo. 13 Microtubules in Sertoli cells In Sertoli cells, microtubules are abundant and are concentrated in a supranuclear region where they extend into apical stalks and processes that support developing spermatids (Christensen 1965; Redenbach and Vogl , 1991). The majority of microtubules are arranged parallel to the longitudinal axis of these cells and have their slow growing or minus ends directed towards the apex of the epithelium (greater than 93%) (Redenbach and Vogl , 1991). This is unlike the situation in some other cell types (such as axons -Bass et al, 1988, Burton and Paige, 1981, Heidemann et al, 1981, melanophores -McNiven et al, 1984, and photoreceptors - Troutt and Burnside 1988) in which microtubules have their positive ends directed towards the periphery of the cell (Figure 4). Experiments from Hermo et al. (1991) indicate that these microtubules have been modified post-translationally since through the use of immunofluorescence they appear to be tyrosinated. This tyrosination may further stabilize the Sertoli cell microtubules, thereby giving the ectoplasmic specializations stable tracts to move along during both the basal and apical translocation of the developing spermatids through the seminiferous epithelium. 14 Precedent for microtubule-based transport and pattern of the Endoplasmic reticulum in General In order for the microtubule-based spermatid translocation hypothesis to be true, molecular motors are hypothesized to link the Sertoli cell microtubules to the endoplasmic reticulum of the ectoplasmic specialization. The general concept of endoplasmic reticulum-microtubule linkages comes from various work indicating associations of the endoplasmic reticulum and microtubules. The endoplasmic reticulum has been known for some time to be closely associated with microtubules. Through electron microscopy Franke (1971) identified crossbridges linking the endoplasmic reticulum to microtubules in cells. Later studies used experimental depolymerization of microtubules to determine if the endoplasmic reticulum was linked to microtubules (Teresaki et al., 1986). These studies showed that the depolymerization of microtubules disrupted the endoplasmic reticulum network in the long term (2 hours), and led to a slow retraction of the endoplasmic reticulum network towards the cell center, indicating that the extended state of the entire endoplasmic reticulum network requires a microtubule system (Teresaki et a l , 1986). In addition to this, recent work done by Klopfenstein et al. (1998) showed that P63 (a 63 kDa non-glycosylated type II integral endoplasmic reticulum membrane protein) binds microtubules in vivo and in vitro and can link the endoplasmic reticulum and microtubules directly (Klopfenstein et al., 1998). This data further corroborates previous work done at the light microscope level which showed that in fixed cells, endoplasmic 15 reticulum tubules colocalized with single microtubules (Teresaki et al., 1986; Dailey and Bridgman 1989). In intact tissues, microtubule-depolymerizing drugs also have been observed to cause drastic reorganization of the endoplasmic reticulum (Vogl et al., 1983). In another study, of cultured cells, where cells were plated in the presence of the microtubule depolymerizing drug nocodazol and allowed to attach and spread overnight, the endoplasmic reticulum did not extend into the thin periphery (Teresaki 1990). However, upon nocodazol removal, many close alignments of endoplasmic reticulum and newly polymerized microtubules were found (Teresaki 1990). These effects of microtubule depolymerization are further evidence that microtubules and the endoplasmic reticulum are closely related (Teresaki et al., 1986). In 1989, Lee et al. found that under no circumstances did the endoplasmic reticulum reform without microtubules. This behavior indicates that in these cells, extension of an endoplasmic reticulum tubule requires a close, physical interaction with a microtubule (Teresaki 1990). During construction, the endoplasmic reticulum forms an intricate polygonal network of membrane tubules and lamellae, which extend throughout the cell (Allan 1995). The branching and intersecting of endoplasmic reticulum tubules to form a network has been hypothesized to be microtubule-dependant, thereby producing the endoplasmic reticulum's characteristic reticular structure (Dabora and Sheetz, 1988; Lee and Chen, 1988; Vale and Hotani, 1988). The extensive evidence of microtubule-endoplasmic 16 reticulum interactions, especially their codistribution and structural interdependence in living cells (Teresaki et al., 1986), is consistent with this hypothesis. Microtubule dependant movements of vesicular organelles are a means for intracellular transport in many kinds of eukaryotic cells (Schnapp et al., 1985; Allan and Kreis, 1986; Kooce and Schliwa, 1985; Kachar et al., 1987; Matteoni and Kreis, 1987; Cole et al., 1993; Hendson et al., 1995; Yamazaki et al., 1996; Criswell and Asai 1998), and microtubule-based motility may be responsible for the formation and movement of the endoplasmic reticulum network as well (Dabora and Sheetz 1988). The results from Lee et al., (1989) suggest that the formation of the microtubule network occurs before the construction of the endoplasmic reticulum, and the movement of the endoplasmic reticulum occurs along the tracts of already formed microtubules. Fluorescent and video microscopy have given results that indicate that motors such as kinesin, cytoplasmic dynein, and myosin are associated with the endoplasmic reticulum (Vale et al., 1985a; Lane et al., 1999). Kinesin, a microtubule-based motility protein that moves towards the plus ends of microtubules at a rate of approximately 1 um/sec (Vale et al., 1985a,b; Howard, 1997; Schnitzer and Block, 1997; Coy et al., 1999), attaches to the endoplasmic reticulum and extends a membrane tubule as it translocates along a microtubule towards the periphery of the cell (Vale and Hotani 1988; Klopfenstein et al., 1998). Conclusion In cells in general there is evidence that microtubule linking and motor proteins are present on the endoplasmic reticulum. 17 Endoplasmic reticulum imaging In order to confirm that any motility events observed in my motility system are actually occurring on an intact ectoplasmic specialization, it would be useful to visualize the endoplasmic reticulum component of the junction plaque. The endoplasmic reticulum can be identified easily in electron micrographs by its morphology, but unfortunately it is more difficult to visualize at the light microscope level. The short chain (6 carbons or shorter) dicarbocyanine dye D i O C 6 (3) has been useful for endoplasmic reticulum identification because it produces a very bright signal, bleaches relatively slowly (Teresaki et al., 1984) and has been used previously to stain the endoplasmic reticulum of the ectoplasmic specialization (Beach and Vogl , 1999). It is likely that the shorter carbon chains of dyes such as D iOC6 (3) do not anchor the dyes in the membrane bilayer as strongly as the longer chain dyes, so that these dyes can permeate across the plasma membrane and associate with the bilayer portion of the internal membranes (Teresaki and Reese 1992). The properties of D i O C 6 and related dyes, as well as their staining characteristics, indicate that they do not associate with components specific to particular membranes, but instead associate with the bilayer portion that is common to biological membranes (Teresaki and Reese 1992); therefore, and unfortunately, D iOC6 labels many membrane organelle compartments (such as mitochondria) in addition to the endoplasmic reticulum (Teresaki et al., 1984; Teresaki and Reese 1992). Even though D iOC6 does not specifically stain the endoplasmic reticulum, it has been used successfully to localize the endoplasmic reticulum network 18 due to the differences in the morphology of it as compared to the other intracellular membranes (Teresaki and Reese 1992). Although there are antibodies to specific endoplasmic reticulum proteins such as BiP and C R T , these have not proven useful for labeling the endoplasmic reticulum of ectoplasmic specializations. Also, antibodies are more difficult than D iOC6 to use in my in vitro system. 19 Motors in general Molecular motors are enzymatic protein machines (mechanoenzymes), whose directed movement along cytoskeletal filaments are driven by the hydrolysis A T P (Howard 1997). These enzymes convert chemical energy from this A T P hydrolysis to mechanical energy in order to generate force for movement. These motors usually associate with specific aspects of the cytoskeleton for example the actin-dependent motors (myosins) and the microtubule-dependent motors (kinesins and dyneins). Many aspects of microtubule-based motility in cells can be attributed to the action of kinesins and dyneins (Valleeand Sheetz 1996). In the case of the microtubule-based spermatid translocation hypothesis, these microtubule-dependent motors are the components predicted to move the developing spermatids from the apex of the Sertoli cell cytoplasm towards the base of the Sertoli cell (kinesin) and then return the spermatids to the apex of the seminiferous epithelium (dynein) for their eventual release into the lumen of the seminiferous tubule. 20 Kinesin Kinesins are microtubule-activated ATPases (adenosine triphosphotases) that are able to move cargo along microtubules mainly, but not exclusively towards the plus end (Vale, 1987). If the microtubule-based spermatid translocation hypothesis is true, then a kinesin-like motor would move spermatids towards the base of the seminiferous epithelium. Conventional (classical) kinesin (~380 kD) consists of two 120 kD kinesin heavy chains and two 64 kD kinesin light chains, and has a total length of 80nm (Vale et al., 1985; Brady 1985; Schnapp et al., 1985; Bloom et al., 1988; Vallee and Sheetz 1996). It has a rod-like structure composed of two, globular heads (10 nm in diameter) positioned at the ends of the kinesin heavy chains (Hirokawa et al., 1989; Scholey et al., 1989), and a long fan-like a-helical coiled coil tail which is associated with the two light chains (Hirokawa et al., 1989). The globular heads bind to the microtubules (and have conserved sequences present throughout the kinesin group of motors), and the tail binds to the organelle cargo (Vikstrom et al., 1992; Okabe et al., 1993; Goldman et al., 1996). The heads also regulate the motor activity (Vallee and Sheetz 1996; Howard 1997), and a stalk (or neck) connects the heads to the tail. The COOH-terminal of kinesin heavy chain is thought to bind membranous cargoes, and this binding could be modulated by the kinesin light chains (Stenoien and Brady 1997; Bloom et al., 1993; Skoufias et al., 1994). Evidence for this comes from experiments showing that antibodies to kinesin light chains, especially to tandem repeats, inhibit fast 21 axonal transport in the axoplasm and release kinesin from purified membrane vesicles in vitro; thereby supporting the idea that the kinesin light chains do in fact play a role in the interaction of kinesin with membranes (Stenoien and Brady 1997). Kinesin's path along the microtubule surface appears to follow one of the protofilaments (Howard, 1997). Since there is one binding site per tubulin dimer, the path distance is 8nm, the dimer's length (Howard, 1997). The actual movement of the kinesin globular heads is thought to occur in a "walking" type of movement with the release of one of the heads only after the other has attached to the next dimer. It is capable of moving along a microtubule for a relatively long distance without the aid of other microtubule-associated proteins (a mechanism of movement termed processive). Intact kinesin has a very low microtubule-activated ATPase activity that is increased dramatically when the molecule is bound to cargo (such as on the surface of anionic beads) (Jiang and Sheetz 199.5). When moving, kinesin usually takes about 100 steps before releasing from the microtubule and each of these steps uses approximately 1 molecule of A T P (Schnitzer and Block, 1997; Hua et al., 1997). Interestingly, constructs of the kinesin molecule without it's tail will support motility and have a high microtubule-activated ATPase activity because there is no inhibitory domain present (Sadhu and Taylor 1992; Hackney, 1994; Gilbert et al., 1995). Antibody labeling studies in sea urchin spindles have localized sub-populations of kinesin to a variety of subcellular organelles, including the Golgi , the endoplasmic 22 reticulum-Golgi intermediate compartment, and other membranes (Schmitz et al., 1994; Skoufias et al., 1994; Lippincott-Schwartz et al., 1995). These sub-populations are termed kinesin-like proteins. Whether or not a protein is a conventional kinesin or is a kinesin-like protein is determined by the sequence of its motor domain (or head) which is about 350 amino acids in length and 4.5 X 4.5 X 7.0 nm in size; this domain is the actual area that hydrolyzes adenosine triphosphate (ATP) and catalyzes movement (Vallee and Sheetz, 1996). On the basis of sequence similarities, the kinesin-like proteins have been grouped into eight major subfamilies (Moore and Endow, 1996). Kinesin proteins have now been grouped into three major superfamilies (Hirokawa, 1998) . Over 100 kinesins have been identified (Sakowicz et al., 1998) and grouped according to the position of their motor domain: NH2-terminal motor domain type, middle motor domain type, and COOH-terminal motor domain type (also referred as N-type, M-type and C-type respectively) (Hirokawa, 1998). Numerous reviews have been published on the various motors and their specific characteristics (Vale and Fletterick, 1997; Block, 1998a,b; Hirokawa, 1998; Hirokawa et al., 1998; Mandelkow and Hoenger, 1999) . 23 Dynein If the microtubule-based spermatid translocation hypothesis is true, then a molecular motor would move developing spermatids from the base of the seminiferous epithelium to the apex. Although a few members of the kinesin family are capable of moving towards the minus ends of microtubules, cytoplasmic dynein is a more likely candidate for this task. There are two main reasons for this, 1) to date there have not been any minus-end directed motors found in mammals and 2) the 74 kD intermediate chain of dynein has been immunolocalized to an area consistent with the location of the ectoplasmic specializations in rat testes (Miller et al., 1999). Cytoplasmic dynein (discovered in 1987 and previously referred to as microtubule-associated protein IC (MAP1C)), is a brain cytoplasmic form of the ciliary and flagellar ATPase, dynein (Vallee et al., 1988). This member of the dynein superfamily is distinguished from axonemal dyneins by it's biochemical characteristics, such as, stimulation of ATPase activity by different concentrations of microtubules, by it's subunit composition and by it's substrate specificity (Collins and Vallee, 1989). Conventional cytoplasmic dynein is a massive retrograde moving (towards microtubule minus ends) multisubunit complex (1.2 M D ) composed of two heavy chains (~530 kD), three intermediate chains (74 kD), and four light intermediate chains (~55 kD) (Pashal et al., 1987; Lye et al., 1987; Shroer et al., 1989; Shnapp and Reese, 1989; Holzbauer and Vallee 1994; Schroer, 1994). The cytoplasmic dynein heavy chain consists of 4644 amino acids in rat and contain potential sites of nucleotidase activity (Gibbons et al., 24 1991;Ogawa, 1991; L i etal., 1993; L i et al., 1994; Lye etal., 1995) at phosphate binding pockets (P-loops) in its central region (Holzbauer et al., 1994; Schroer, 1994; Koonce et al., 1992; Mikami et al., 1993; Zhang et a l , 1993; Eshel et al,. 1993). Ten to fifteen dynein heavy chains (the majority of which are likely ciliary or flagellar) have already been identified in each of several organisms, including rat (Tanaka et al., 1995; Vaughan and Vallee, 1995), sea urchin (Gibbons et al., 1994), Drosophila (Rasmusson et al., 1994), Chlamydomonas (Goodenough et al., 1987; Piperno et al., 1990; Wilkerson et al., 1994), and humans (Vaughan et al., 1995). Cytoplasmic dynein interacts with many different organelles. In vitro analysis of the interaction of cytoplasmic dynein with membranes (Yu et al., 1992; Fath et al., 1994; Ferro and Collins, 1995) and studies of broken cell preparations (Aniento et al., 1993; Corthesy-Theulaz et al., 1992; Lafont et al., 1994) have supported an interaction with endosomes, apical exocitotic vesicles (in epithelial cells), component membranes of the Golgi apparatus, synaptic vesicles (Lacey and Haimo, 1992) and membranes of the endoplasmic reticulum (Allan, 1995; Y u et al., 1992). In the case of the endoplasmic reticulum interaction, phosphorylation has been shown to regulate endoplasmic reticulum movement by controlling the activity of cytoplasmic dynein bound to the endoplasmic reticulum membrane (Allan, 1995). In addition to intermediate and light intermediate chains, cytoplasmic dynein is associated with the 10 subunit protein complex dynactin (Gill et al., 1991) which also is 25 thought to be the membrane receptor for dynein and which is reported to activate the motility of membrane-bound dynein (Schafer et al., 1994). 26 Multiple motors With the identification of so many molecular motors, there are increasing instances where multiple motors are expressed simultaneously in a cell (germ cell manchette and germ cell residual cytoplasm, Hall et al., 1992; axons, Muresan et al., 1996; cultured NIH-3T3 fibroblasts, N R K fibroblasts, BHK-21 fibroblasts and rat glial cells, Lin et al., 1996). One possible reason for the presence of multiple motors is that they are required for multiple transport phenomena that occur within the same cell (Vallee and Sheetz, 1996). This transport is determined by 1) the polarity of the microtubules, 2) the type of motors present, 3) the temporal recruitment of the motors, 4) the targeting of the motors to specific sites in cells and 5) the regulation of the motors. In the spermatid transportation system it is likely that these motor proteins will take advantage of similar elements as those found in other systems in order to move the developing spermatids from the apex of the seminiferous epithelium to the base (with one type of motor) and from the base of the epithelium to the apex (with another type of motor). Since the 74 kD intermediate chain of dynein has been found at ectoplasmic specializations (Miller et al., 1999) and to date neither a plus-end directed dynein nor a single motor which can move in both the plus and minus-end directions has been identified, it is likely that multiple motors are at work in translocating the spermatids during spermiogenesis. 27 Motors in Testis Kinesin in Testis The kinesin superfamily of molecular motors comprises proteins that participate in a wide variety of motile events within the cell (Sperry and Zhao, 1996). There is now evidence for a number of kinesins in testis (Sperry and Zhao, 1996). The 85K and 115K subunits of kinesin II are present in echinoderm sperm and through immunofluorescence these proteins have been localized to the sperm flagella and midpiece (Henson et a l , 1997). Kinesin II subunits are related to K IF3A and KIF3B from mouse. These KIFs appear to be coordinately expressed, with the highest levels of KIF3 transcripts and polypeptides being detected in brain and brain plus testis samples (as shown by western blots), respectively (Aizawa et al., 1992; Kondo et al., 1994; Yamazaki et al., 1995a,b). Through these findings Henson et al. (1997) hypothesize that kinesin II in sperm may play functional roles in interflagellar transport and/or the formation of flagella during spermatogenesis. Recent immunofluorescent evidence from Miller et al. (1999) shows kinesin II present at the flagella of rat spermatids, further corroborating this claim. Members of the kinesin superfamily share a highly homologous head domain responsible for force generation, which is attached to a divergent tail domain, which is thought to couple the motor domain to its target cargo (Sperry and Zhao, 1996). Sperry and Zhao (1996) identified a group of kinesin-related proteins from rat testes (termed testes KRP1 through KPR6) that may define new kinesin subfamilies. KRPs (kinesin related 28 proteins) contain a mechanochemical domain that is 30-40% identical to other family members but this sequence conservation does not extend to the remainder of the molecule (Sperry and Zhao, 1996). These variable regions, often termed tail domains, are thought to specify the target and cellular function of each K R P (Sperry and Zhao, 1996). Rat testes contain at least six kinesin-related proteins, three of which are expressed exclusively in the seminiferous epithelium (Sperry and Zhao, 1996). Five of the six testes KRPs identified are expressed primarily in testes (as demonstrated through K R P R N A slot blot analysis of gene expression) (Sperry and Zhao, 1996) and three of these (KRP2, KRP3 , and KRP5) appear to be expressed primarily in the meiotically active cells of the seminiferous epithelium (as was seen through the in situ localization of the 35 K R P gene expression through the dark and light field viewing of S-labeled R N A probes that were complimentary to the K R P mRNAs). The other two, KRP4 and KPR6, show a strikingly different distribution, being expressed at lower levels than K R P 2 , KRP3 , and KRP5 , and being in a different functional compartment of the epithelium (Sperry and Zhao, 1996). Two testes KRPs, KRP2 and KRP6 , are related to the already identified superfamily members KIF2 and Eg5 respectively, but the remaining KRPs do not match any known K R P and may represent motors with previously unknown functions (Sperry and Zhao, 1996). These new KRPs are candidates for testes-specific motors with roles in movements unique to spermatogenesis and spermiogenesis including chromosome 29 segregation, nuclear shaping of the sperm head, flagellar movements, and transport of spermatozoa within Sertoli cell crypts (Sperry and Zhao, 1996). KRP2 is a new member of an emerging kinesin subfamily of central domain KRPs and unlike KIF2, KRP2 is expressed only in the testes (Aizawa et al., 1992; Sperry and Zhao, 1996). Worth noting is that KRP3, due to its identity to a flagellar K R P , is a good candidate for a motor involved in the formation and functioning of motile sperm (Sperry and Zhao, 1996). In addition to this, immunoreactive cytoplasmic dynein and kinesin have been detected in rat testes and showed a stage-dependent distribution (Hall et al., 1992). The latter authors expected no kinesin staining in the Sertoli cell cytoplasm or at ectoplasmic specializations but found kinesin at the germ cell manchette from stages X-V I and at the germ cell residual cytoplasm from stages V-VI (Hall et al., 1992). The dynein staining was present in the Sertoli cell cytoplasm through out spermatogenesis and at ectoplasmic specializations from stages IX to X IV as well as being present at the germ cell manchette at stages I-IV and in the residual germ cell cytoplasm at stages V-VI (Hall et al., 1992). Kinesin in Sertoli cells Thus far, there is very little evidence for a kinesin in Sertoli cells, other than for the localization of the SUK4 (anti-sea urchin kinesin) monoclonal antibody to the Sertoli cell trans-Golgi network (Johnson et al., 1996; Ingold et al., 1988). There is no evidence for a kinesin at ectoplasmic specializations. 30 Dynein in testis Rat testis has proven to be a rich source of cytoplasmic dynein (Collins and Vallee, 1989; Neely et al.,1990; Hall et al., 1992). The yield of dynein from testis is approximately 70 pg/g of tissue, making this the best source of cytoplasmic dynein of all tissues examined thus far (Collins and Vallee, 1989). This yield is almost three times higher than that in brain (Collins and Vallee, 1989). Two of the identified mammalian cytoplasmic dynein heavy chains are D H C l a and D H C l b (Criswell and Asai, 1998). D H C l a is conventional cytoplasmic dynein and is found in all organisms that have been examined thus far (Criswell and Asai, 1998). D H C l b is expressed in organisms that have multiple dyneins, and has been implicated in the intracellular trafficking of molecules in unciliated and ciliated cells (Criswell and Asai, 1998). In the case of the testis, cytoplasmic dynein has been found to contain a large amount of dynein heavy chain that is immunoreactive with antibodies raised against a peptide sequence derived from D H C l b (Criswell and Asai, 1998), and sucrose gradient centrifugation and V I photolysis identified four separate cytoplasmic dyneins in rat testis: D H C l a , D H C l b , and two other isoforms (Criswell and Asai, 1998). Through these experiments, Criswell and Asai (1998) concluded that rat testis contains three D H C l b - l i k e proteins, and one of these is the product of the D H C l b / D H C 2 gene, implying that there are likely multiple dyneins present to perform different tasks. This conventional cytoplasmic dynein, herein called D H C l a , is present in all eukaryotes and forms a two-headed homodimer (Vallee et a l , 1988; Neely et al., 1990). In both 31 unciliated and ciliated mammalian cells, there are multiple cytoplasmic dynein heavy chain isoforms that are speculated to perform separate tasks (Criswell and Asai, 1998). D H C 1 , which is the same as D H C l a , is conventional cytoplasmic dynein; DHC2 , which is the same as D H C l b , is involved in positioning the Golgi apparatus; and D H C 3 is associated with what may be transport intermediates between intracellular compartments (Vaisberg et al., 1996). 32 Dynein in Sertoli Cells The rat seminiferous epithelium has fourteen morphologically distinct stages, culminating in the release of mature sperm at the end of stage VIII (Leblond and Clermont 1952). Cytoplasmic dynein (MAP1C, Neely et al., 1990) was identified and immunolocalized to the Sertoli cell cytoplasm during all stages of spermatogenesis (Hall et al., 1992; Neely and Boekelheide, 1988), with a peak in the apical cytoplasm during stages IX-XIV (Hall et al., 1992). It has recently been localized, using an antibody to the 74kD intermediate chain, to areas of the Sertoli cell associated with spermatid translocation and known to contain ectoplasmic specializations (Miller et al., 1999). This is an important finding since it would be consistent with the hypothesis of motors being present on the cytoplasmic side of the endoplasmic reticulum on the ectoplasmic specialization. This could account for the movement of the developing spermatids from the Sertoli cell nucleus to the apex of the Sertoli cell and would be consistent with the microtubule translocation hypothesis. The motor has yet to be localized using immunoelectron microscopy to the endoplasmic reticulum component of the junction plaque. 33 Dynein in Spermatogenic Cells Dynein is associated with the microtubular manchette and the nuclear envelope in the spermiogenic steps 7 through 14 spermatids, with strong expression in steps 8 to 12 (Yoshida et al., 1994). Dynein is concentrated in the region between the nuclear envelope and the closest microtubules (Yoshida et al., 1994). These observations indicate that, during spermiogenesis, the microtubular manchette and the nucleus might assemble a mechanical structure that is driven by cytoplasmic dynein which may function in reshaping the nucleus (Yoshida et al., 1994). 34 The Microtuhule-Based Spermatid Translocation Hypothesis and the Thesis Problem The microtubule-based spermatid translocation hypothesis can be stated as follows: Molecular motors (or motor proteins), that link microtubules to ectoplasmic specializations, generate the force for the moving spermatids throughout the seminiferous epithelium (Vogl et al., 1991). The motor proteins are anchored to the cytoplasmic face of the endoplasmic reticulum component of the junction plaque and the microtubules act as tracts along which spermatids are moved from the apex of the Sertoli cell towards the Sertoli cell nucleus (stages III-V) and back to the apex (stage VI and VII) for eventual release into the lumen of the seminiferous tubule (Figure 5). The occurrence of translocation both in a basal and an apical direction indicates that there may be at least two motors involved in generating the movement. In other systems, microtubule-based motors (or mechanoenzymes) consist of the cytoplasmic kinesins and dyneins. Evidence consistent with the microtubule-based spermatid translocation hypothesis consists of the following: (1) There is an abundance of motor proteins in the testis (Sperry and Zhao, 1996; Yamazaki et al., 1996; Henson et al., 1997; Criswell and Asai, 1998; Miller et al., 1999), (2) Microtubules are oriented parallel to the direction of spermatid movement (Fawcett, 1975), (3) Linkages occur between the endoplasmic reticulum of the ectoplasmic specialization and microtubules (Russell, 1977), (4) Decreased numbers of microtubules are bound to the junction plaque after the addition of A T P (likely due to the cycling off of the microtubules as was shown from binding assay experimentation) (Vogl, 1996), (5) Immunolocalization of the 74kD intermediate chain 35 of cytoplasmic dynein at ectoplasmic specialization (Miller et al., 1999), (6) Isolated ectoplasmic specializations support microtubule movement (Beach and Vogl ,1999). 36 T h e s i s P r o b l e m (H y p o t h e s i s ) In the g e ne r a l m o d e l o f m i c r o t u b u l e - b a s e d s p e rm a t i d t r a n s l o c a t i o n d e s c r i b e d a b o v e , e c t o p l a s m i c s p e c i a l i z a t i o n s m o v e a l o n g m i c r o t u b u l e s i n b o t h d i r e c t i o n s in the S e r t o l i c e l l . In o t he r w o r d s , the j u n c t i o n p l a q u e s m o v e f i r s t t owa r d s the p l u s ( fas t g r o w i n g ; b a s a l l y p o s i t i o n e d i n v i v o ) a n d t h e n t o w a r d s the m i n u s ( s l o w g r o w i n g ; a p i c a l l y p o s i t i o n e d i n v i v o ) ends o f the Se r t o l i c e l l m i c r o t u b u l e s d u r i n g s pe rmatogene s i s . I f m o v e m e n t o f the j u n c t i o n p l a q u e is m i c r o t u b u l e - b a s e d a n d d o e s o c c u r i n b o t h d i r e c t i o n s , t h e n w h e n e x o g e n o u s m i c r o t u b u l e s are a d d e d to i s o l a t e d e c t o p l a s m i c s p e c i a l i z a t i o n s in vitro, I s h o u l d be ab l e to detect t w o p o p u l a t i o n s o f m i c r o t u b u l e s -t h o s e m o v i n g w i t h the i r p l u s e nd s f o r w a r d ( dyne i n- l i k e ) a n d those m o v i n g w i t h the i r m i n u s ends f o r w a r d (k i nes i n- l i ke ) ( F i g u r e 6). T o v e r i f y the p r e d i c t i o n that the j u n c t i o n p l a q ues s uppo r t m i c r o t u b u l e m o v e m e n t in t w o d i r e c t i o n s , I e s t a b l i s h e d an i n v i t r o m o t i l i t y a s s a y u s i n g p o l a r i t y m a r k e d m i c r o t u b u l e s ( d e s c r i b e d i n deta i l i n the ma te r i a l s a nd me t h o d s sect i on ) . T h e a s say b a s i c a l l y c o n s i s t e d o f c o n s t r u c t i n g p o l a r i t y -m a r k e d m i c r o t u b u l e s , a d d i n g t h e m to i s o l a t e d e c t o p l a s m i c s p e c i a l i z a t i o n s a n d a s s a y i n g t h em f o r mo t i l i t y ( F i g u r e 3). 37 Figure 1 . Diagram of a section of the seminiferous epithelium showing a typical Sertoli cell, some of it's organelles and it's embedded germ cells. This picture was modified from Fawcett(1975). 38 31 F i g u r e 2. D i a g r a m m a t i c r e p r e s e n t a t i o n o f t h e m i c r o t u b u l e t r a n s l o c a t i o n h y p o t h e s i s ( g i f t f r o m A . W . V o g l ) . T h e s p e r m a t i d m o v e s f r o m t h e a p e x o f t h e S e r t o l i c e l l t o w a r d s t h e b a s e o f t h e S e r t o l i c e l l a n d r e t u r n s to t h e a p e x o f t h e S e r t o l i c e l l a l o n g m i c r o t u b u l e s t h a t a c t as t r a c t s o n w h i c h m o l e c u l a r m o t o r s r i d e w h i l e b e i n g a t t a c h e d t o t h e e c t o p l a s m i c s p e c i a l i z a t i o n . 40 Spermatid 41 Figure 3. Diagrammatic representation of the experimental model. The ectoplasmic specialization which (moving cytoplasmically) begins at the Sertoli cell plasma membrane followed by the hexagonally packed actin bundles and the endoplasmic reticulum. Attached to the cytoplasmic side of the endoplasmic reticulum are hypothesized to be anterograde and retrograde moving motors (possibly cytoplasmic dynein and kinesin) and attached to the motors is a G M P C P P polarity marked microtubule with its brightly labeled rhodamine tubulin seed and its long, positive, dimly rhodamine tubulin labeled tail and short, negative, dimly rhodamine tubulin labeled end. 42 Molecular Motors Sertoli Cell Plasma Spermatid Plasma Polar Microtubule I Endoplasmic Reticulum Membrane Membrane Minus E n d " Seed Actin o d o o O O O Q O O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o O O O O o o o o o o o o o o o o o o O O O O o o o o o O O O O O O O O O O O O O o o o o o O O O O o o o o o O O O O o o o o o 43 Figure 4. Diagrammatic representation of microtubule polarity in a cultured cell, a neuron and a Sertoli cell. Note the reverse polarity of the microtubules in the Sertoli cell. This picture is a modified gift from Dr. A . Wayne Vogl. 44 Figure 5. Four paneled diagrammatic representation of the microtubule translocation hypothesis from Vogl et al., (1991). The spermatid moves from the apex of the Sertoli cell towards the base of the Sertoli cell and returns to the apex of the Sertoli cell along microtubules that act as tracts on which molecular motors ride while being attached to the ectoplasmic specialization. 46 Figure 6. Direction of movement of both anterograde and retrograde molecular motors depicted as the kinesin and dynein motors respectively. The anterograde motor moves cargo towards the (+) end of the microtubule. When the motor is anchored in position, as it is in my in vitro assay system, the polar microtubule moves leading by the brightly labeled seed. The retrograde motors move towards the (-) ends of the microtubules. When these motors are anchored, microtubules move trailing with the seed. 48 P l u s E n d M o v e m e n t s (eg . K i nes i n ) Microtubule Anchored Motor Anchored Materials and Methods Animals A l l animals used in this study were Sprague-Dawley rats, ranging in weight from 255 grams to 604 grams. They were obtained from a colony maintained at the University of British Columbia animal care facility and then were held in a room in the Department of Anatomy until use. Chemicals and Reagents Unless otherwise indicated, all chemicals and reagents were obtained from the Sigma Chemical Company (St. Louis, M O / U S A ) . The bovine brain tubulin and rhodamine labeled tubulin were purchased from the Cytoskeleton Company (Denver, C O / U S A ) at a stock concentration of lOmg/ml. The Paraformaldehyde was obtained from Fisher Scientific (Vancouver, B C , Canada). Polar Microtubule Construction Three methods of polar microtubule construction were attempted. O f the three only the final method was routinely used for the motility assay. The three methods were the "Cytoskeleton" method, the " N E M " method and the " G M P C P P " method. Each is described in the following sections. 50 Cytoskeleton Method of Polar Microtubule Construction The Cytoskeleton method of polar microtubule construction was recommended by Cytoskeleton and came with reagents purchased together as a kit from this company. This kit contained all of the necessary components to produce polarity labeled microtubules. The protocol consisted of preparing taxol stabilized microtubule seeds constructed by polymerizing microtubules from a pool of tubulin containing a high concentration of rhodamine labeled tubulin and polymerizing ends off of the brightly labeled seeds from a pool of tubulin containing a low ratio of labeled to unlabelled tubulin. To do this 10 u.1 of 2mM taxol (in dry D M S O ) was added to 1 ml of microtubule polymerization buffer consisting of 80mM PIPES (pH 6.8), ImM E G T A and ImM MgC12. This was called the T P B solution and was kept at room temperature. To grow the seeds, 60 pi of the T P B solution was added to 2 pi of 10 mg/ml "seed" rhodamine tubulin. After thoroughly mixing this and allowing this tube to stay at room temperature for 5 minutes, the seeds should be 1 urn. At that time they were placed on ice where they could stay for up to thirty minutes. To grow the dimly labeled microtubule ends, 70 pi of microtubule polymerization buffer was added to 2 u.1 of dimly labeled rhodamine tubulin at 10 mg/ml. This was incubated at 37 °C for 5 minutes. After the incubation, 1 u.1 of seeds were added to the tube and the solution was incubated for exactly 5 minutes. Once completed, 240 u.1 of the T P B solution was added to the tube to stop the polymerization of the dimly labeled tubulin 51 ends onto the brightly labeled seeds. At this point a stable population of polarity marked microtubules labeled with the seed at their minus ends should be constructed. Monomeric tubulin was removed by layering 150 u.1 of the polar microtubules onto a 800 pi cushion buffer consisting of 80mM PIPES (pH 6.8), ImM E G T A , ImM MgC12, 30% glycerol and 8 ul of 2 m M taxol (in dry D M S O ) followed by centrifugation at 100,000g at 25 °C for 30 minutes. This should produce a pellet of microtubules with which the first 200 pi of solution was discarded. At this point the cushion/microtubule interface was washed twice with 100 pi of the T P B solution and then the remaining cushion buffer was discarded and the pellet was gently resuspended in 100 pi of T P B solution. 52 N E M method of Polar Microtubule Construction The N E M (N-ethylmaleimide) method of polar microtubule construction was that published by Scholey (1993). Widely used because it produces polar microtubules of unambiguous polarity, since N E M prevents polymerization from the minus-end. This method of building polar microtubules takes approximately 7 hours. The seeds take 6 hours to construct and the dimly labeled microtubule positive end takes another hour. Although this method was more successful than the pervious, it still was unacceptable (due to it's inconsistency in polymerizing polar microtubules arid its extremely low yield) for use in the motility assay even after extensive modification. The original N E M method of polar microtubule construction is as follows. This method required the construction of glycerol seeds followed by the construction of rhodamine labeled seeds. Once completed the dimly rhodamine labeled microtubule positive end only is grown off of the brightly rhodamine labeled seed. The first part of growing N E M polar microtubules consists of building glycerol microtubules, which were sheared to form nucleating agents. These agents acted as centers for the polymerization of brightly labeled rhodamine tubulin, which in turn were fragmented to form the rhodamine labeled seeds during the future steps of this method of polar microtubule construction. To build the glycerol microtubules, 50 pi of glycerol, 0.5 pi of 1.0 M MgC12 and 1.5 pi of . ImM GTP were added to 100 pi of unlabeled tubulin at 4mg/ml. This was well mixed and incubated at 37 °C for 50 minutes. Once completed the glycerol microtubules were sheared 5 times with a 26 gauge needle to 53 make short microtubule fragments - the glycerol seeds. The next step involved diluting the glycerol seeds into rhodamine labeled tubulin at 20 uM. This was accomplished by mixing on ice, 8.33 pi of 4 mg/ml unlabeled tubulin, 10 u.1 of 10 mg/ml rhodamine labeled tubulin, 15 pi of BRB80 (80mM PIPES and ImM MgC12 (pH 6.8)) and 0.33 ul of lOOmM GTP. This was warmed at 37 °C for 1 minute at which point 1 pi of glycerol seeds were added to the mixture, well mixed and incubated at 37 °C for another 60 minutes. After the 60 minute incubation, the newly constructed brightly labeled rhodamine microtubules were cross-linked through the addition of 1/10 the volume of pre-warmed 15 m M E G S in D M S O for 60 minutes at 37 °C. Following the cross-linking, 2 ml of BRB80 containing 50% sucrose, lOmM potassium glutamate and 0.1% p-mercaptoethanol were added to the cross-linked microtubules to quench and dilute the solution for 60 minutes at 37 °C. After this incubation the rhodamine labeled microtubules were sheared four times with a 26 gauge needle which were now called the rhodamine labeled seeds. These seeds were cleaned to remove any excess unpolymerized tubulin by diluting the seed mixture 1:1 with BRB80 and layering on a step gradient, 2 ml of seeds, 1 ml of 40% sucrose in BRB80 + 0.1% P-mercaptoethanol, and 2 ml of 75% sucrose in BRB80 + 0.1% P-mercaptoethanol. This was then spun at 37 °C for 40 minutes at 30,000 rpm in a SW 65Ti rotor. The seeds were then collected at the 40%/75% interface. These seeds are reputed to be stable at room temperature for 2 months. Once the seeds were successfully constructed, dimly labeled microtubule positive ends were polymerized onto the brightly labeled seeds. This was accomplished by adding 5 4 13.635 pi of BRB80, 1.3 pi of 50 m M N E M (to get a final concentration o f l m M N E M ) and 0.1 m M G T P to 50 pi of unlabeled tubulin at 5.2 mg/ml to get a final N E M tubulin concentration of 4mg/ml. This was incubated a 0 °C for 10 minutes. Once completed, 8mM P-mercaptoethanol was added to the mixture, well mixed and incubated at 0 °C for 10 minutes to inactivate the excess N E M . Now that the N E M tubulin solution was made the next step was to prepare the dimly labeled rhodamine solution. This was accomplished by adding 45.5 pi of N E M tubulin to 50 pi of unlabelled tubulin at 5.2 mg/ml and 15 pi of BRB80 to get a final tubulin concentration of 4 mg/ml while having an N E M tubulin : tubulin ratio of 0.7 : 1.0. The next step was to make the dimly labeled rhodamine tubulin/NEM tubulin solution, which was used to build the dimly labeled positive ends of the polar microtubules. This was done by mixing 37.5 pi of the 4mg/ml N E M tubulin : tubulin mixture with 2.0 pi of 10 mg/ml rhodamine tubulin, 0.1 pi of 0.1M G T P and 60.4 pi of BRB80. This gave the 10:1 ratio of 1.5 mg/ml N E M tubulin : tubulin to .15 mg/ml rhodamine tubulin needed to make the dimly labeled microtubule positive ends. This mixture was warmed at 37°C for 1 minute. After the 1 minute incubation, 10 pi of the previously made seeds were added to the mixture and were incubated at 37 ° C for 30 minutes to grow the long positive microtubule ends. The microtubules then were stabilized by adding 900pl of, pre-warmed to 37 °C , BRB80 containing lOmM taxol. Care was taken to avoid shearing the newly polymerized polar microtubules. Monomeric tubulin was separated from the polymerized microtubules as follows. 10pl of 20mM taxol was added to 1ml of cushion buffer (that was used in the cytoskeleton 55 method of polar microtubule construction). 300 pi of this solution was then added to the base of each of three tubes used in the SW 65Ti rotor. Following this 350 pi of the polar microtubules were gently layered above the cushion buffer in each of the tubes. They were then spun at 100,000g and once completed the polar microtubules were resuspended with 500 pi of PEM/250 containing 5 u.1 of 20mM taxol. The N E M polar microtubules were now rid of the majority of monomeric tubulin. Some examples of changes made to this polar microtubule assembly method, were as follows. During the seed production, the amount of glycerol seeds was increased to try to get more rhodamine labeled seeds and in turn decrease the amount of monomeric tubulin left in solution. The ratios of labeled to unlabeled tubulin were optimized to have the brightest seeds possible without the seeds depolymerizing under the fluorescent beam and without them sticking to the glass slides during the kinesin and testicular cytosol control runs. The non-hydrolyzable analogue of G T P called G M P C P P (used in the following G M P C P P method of polar microtubule construction) was used to replace G T P to stop seeds depolymerization. The amount of E G S was modified to try to also stabilize to seeds. During the cleaning of the seeds the amount of centrifugation time was increased to try to get cleaner seeds. Azide was used at a concentration of .02% to try to stop the degradation of the seeds while being kept at room temperature, since they were supposed to be stable for 2 months but in my case never lasted over 2 days. During the microtubule polymerization onto the seeds the amount of rhodamine to N E M unlabelled tubulin was optimized. In addition to this the microtubule lengths were also optimized. This was done by controlling the incubation times and increasing the volume of BRB80 56 + taxol to 2900 pi which was the minimum volume that would stop the microtubule polymerization. 57 G M P C P P Method of Polar Microtubule Construction This protocol for polarity-marked microtubules proved to be the best of the three protocols tried. The method that was used originally came from Howard and Hyman (1993). It was then modified by Dave Coy and Will Hancock and was finally adjusted by me to produce polarity marked microtubules suitable for use in my motility assay. Seed construction To make the brightly rhodamine labeled seeds, a mixture of labeled and unlabeled tubulin (RTU) was made. R T U is a combination of 10.5 p L of 10 mg/ml tubulin, 4.5 p L of 10 mg/ml rhodamine labeled tubulin and 22.5 p L of P E M (80mM PIPES, ImM MgC12, ImM E G T A , pH 6.9) which gives a final tubulin concentration of 4mg/mL. Once this was made, 32 u M of the R T U tubulin and 0.15 m M of G M P C P P (a nonhydrolyzable analogue of GTP) were combined and were incubated for exactly 30 seconds at 37°C in a water bath. This produced the seeds for constructing the polarity marked microtubules. Polar microtubule construction Prior to making the seeds as described above, 30.8 p L of P E M , 3 p L of R T U tubulin, 3 p L of 9mg/mL tubulin and 1.6 p L of 0.1 M M g C ^ were combined into each of three reaction tubes and placed on ice. Fifteen seconds before the seeds incubated as denoted above, 1.6 p L of 25mM G T P was added to each of the three reaction tubes and the tubes were thoroughly mixed. The reaction tubes were placed in the 37°C water bath to warm-58 up for 1 minute. After the minute long reaction tube incubations, 1.3 pL of the seeds (that were grown for 30 seconds during the 1 minute reaction tube incubation) were added to each of the reaction tubes and well mixed. The reaction tubes were then placed back in the 37°C water bath for 4 minutes to grow microtubule ends off of the brightly labeled seeds. These polymerizing microtubule ends grew off of the positive and negative ends of the seeds but the microtubule polarity was clearly visible at the negative ends of the microtubules. After the three minute incubations, the polar microtubules were stabilized by diluting all of the polarity marked microtubules from each tube (via cut-tipped pipette) into three pre-warmed tubes containing 360 uL of PEM containing .OlmM taxol. To clean the polar microtubules, each tube containing the polar microtubules was spun through 100 uL of PEM/250 (80mM PIPES, ImM MgCl 2 , ImM E G T A , 250 mM sucrose, 10 ug/ml soybean trypsin inhibitor, 0.5 ug/ml Leupeptin, 0.5 uL/ml Pepstatin and .ImM PMSF) containing .01 mM taxol and 200 uL of PEM/60% at the base of each tube, at 30,000 RPM for 10 minutes using the SW65 Ti rotor. Once spun down, 400 pL of the PEM cytosol was removed from each tube and the remaining microtubules found within the 100 pi of PEM/250 were carefully removed, pooled, aliquoted, then frozen in liquid nitrogen and placed in a -70 freezer (freezing the aliquots was suggested by Will Hancock and John Howard and is done routinely in their laboratory). One aliquot was left out to check for microtubule polymerization and on subsequent days, the microtubules were simply thawed when needed. 59 I considered a microtubule "polarity labeled" if there was either a seed flanked by two ends of varying length (one long and one short) or, if there was only one end attached to a seed, then that end must be visibly longer than the seed. Both of these criteria were used in assessing if these microtubules were polar. If there was any doubt then the microtubule was considered non-polar. 60 i Preparation of the testicular cytosol The preparation of the testicular cytosol was done using 12 testes from 6 Spraugue-Dawley rats. First the SI cytosol was isolated, then the SI A cytosol was made from the SI cytosol following a protocol similar to that of Beach and Vogl (1999). Once under deep anesthesia, the testes from the 6 rats were removed. Each testis was perfused with PBS (150mM N a C l , 5mM KC1, 3.2mM Na2HP04, .8mM K H 2 P 0 4 ; pH7.3), placed in a Petri dish containing fresh ice cold PBS, decapsulated and then minced into small pieces using 2 scalpels. This material was then sedimented for 3-5 minutes in a clinical centrifuge at a low setting of 3. Once sedimented, the final volume of tissue was determined and was re-suspended by adding 90% of the volume of tissue, with homogenization buffer (PEM/250 + l.OmM Dithiothreitol (DTT)). The pellet was then homogenized by passing it through a 10ml Kimble disposable pipette. Following this, the nuclei were removed from the tissue by centrifuging the tissue in a T L A 120.2 rotor at 6,000g for 15 minutes at 4°C. The cytosol was removed and was centrifuged in a T L A 120.2 rotor at 200,000g for 30 minutes at 4°C. The lipid layer was removed and the remaining solution constituted the SI cytosol. To prepare the SI A cytosol (tubulin depleted), taxol and G T P were added to 7mls of the SI cytosol to a final concentration of 20mM and ImM respectively. This was incubated at 33°C for 30 minutes. Once completed, this solution was centrifuged in a T L A 120.2 rotor at 40,000g for 30 minutes at 33°C. The cytosol was then collected. This was the 61 SI A cytosol. Samples were set aside to determine the protein concentration of the SI A cytosol using the Shimadzu ultraviolet-VIS Recording Spectrophotometer (ultraviolet-1 6 0 U ) . Remaining samples were aliquoted and frozen for use in the motility assays. 62 Spermatid and Ectoplasmic specialization isolation Elongate spermatids together with their attached Sertoli cell ectoplasmic specializations (junction plaques) were obtained from the testes of Spraugue-Dawley rats as described by Beach and Vogl (1999). The method can be summarized as follows. Animals were anesthetized and their testes were removed. The organs were placed in a Petri dish and decapsulated. The seminiferous tubule mass was then transferred into a Petri dish containing cold PEM/250 and diced into small pieces using two scalpels. Under a Zeiss dissecting microscope fitted with a dark field condenser, microprobes were used to squeeze the epithelium out of individual seminiferous tubules. The epithelial material was collected using a 37mm capillary 0.3-200 pL microFlex pipette tip and transferred to a conical plastic tube on ice. The material was collected for up to thirty minutes at which time the collected material was sedimented in a clinical centrifuge at a low setting of 2 for 3 minutes. The cytosol was removed and 150 p L of fresh cold PEM/250 buffer was added. The tissue was then gently aspirated via 7 slow, gentle passes using a microFlex pipette to fragment the epithelium. This was then added to three 5% step sucrose gradients (from 60% to 30%) in 0.8 ml ultraclear tubes and was centrifuged in the SW65 T i rotor for exactly 7.5 minutes at 5,000 R P M at 4 °C . Using a 23 gauge needle on a l m L syringe, the material was then collected from the 40%/45% interface by piercing through the tube wall. The fractions from the three tubes were pooled and added to l m L of cold blocking buffer (PEM/250 containing 5 mg/ml casein) for 5 minutes. This was done to block all non-specific sites on the ectoplasmic specializations. The tissue was spun at a low speed of 6 on an Epindorf tabletop centrifuge to loosely pellet the cells and after two minutes of spinning, the tube was rotated 180° and was spun for another two 63 minutes. Once completed the blocking buffer was removed and the cells were resuspended in 50 pi of PEM/250 with protease inhibitors (Pis). These cells were now ready to be used in the motility assay. 64 Preparing the perfusion chamber Using motility chambers purchased from the Cytoskeleton Company (Denver, CO) , 5 uL of the isolated cells with their attached ectoplasmic specializations were added to 10 u L of polar microtubules and allowed to spin on a rotator at room temperature for 10 minutes to allow the polar microtubules to attach to the cells (Figure 7). Following this, the mixture was added very slowly (via cut-tipped pipette) to the chamber for 10 minutes of incubation at room temperature to allow the cells to attach to the slide. When this was completed, 10 p L of wash buffer (PEM+ 2uM Taxol+1 mg/ml casein) was added to the chamber to wash away any unbound cells, ectoplasmic specializations or polar microtubules. 65 The motility assay Once all of the components were added to the motility chambers, the motility assay was initiated. The motility assay was recorded using a Zeiss Axiophot microscope fitted with a C C D digital camera (Princeton Instruments, Trenton, NJ), which was in turn attached to a Power PC. A programmed script using IP Lab Spectrum was started. The script consisted of eighty, three second exposures separated by a three second delay and computer processing time. After 10-16 control frames, 10 p L of warm (33°C) motility buffer (containing A T P ) was added by a hanging drop to the chamber while on the microscope stage and drawn through the chamber via a hanging piece of filter paper (Figure 8) in order to not disturb the very thin polar microtubule focal plane. The script continued until 80 frames were captured and the direction (polarity) of any microtubule movement was determined by animating the captured images. 66 Verification that ectoplasmic specializations were attached to spermatids Once complete, the cell was stained with 10 uL of 100 U/ml coumarin phallacidin (Molecular Probes; Eugene, OR) (in PEM/250) for 5 minutes to visualize the actin found at the ectoplasmic specializations. Following this the cell was stained with 10 uL of 0.5 ug/ml DIOC6 (Molecular Probes; Eugene, OR) (in PEM/250) for 10 seconds to visualize the endoplasmic reticulum cisternae that are also found at the ectoplasmic specializations (Figure 7). These two stains were used to ensure that the ectoplasmic specializations were still intact and that any movement that was documented did in fact occur at the ectoplasmic specializations. 67 Controls For the proper position of the seed Controls for the proper position of the seed in the polar microtubules consisted of either bovine brain or recombinant kinesin (a positive directed motor) being added to the motility chamber in place of the testicular cytosol. The kinesin confirmed that the brightly labeled seeds were positioned at the negative ends of the microtubules by the polar microtubule movement leading by the seed. In addition to this, experiments were also run with both the inclusion and exclusion of 2 mg/ml testicular cytosol present in the motility buffer to ensure that the motility buffer used in the assay could support movement. For the presence of nucleotide To control for the presence of nucleotide (ATP) the first 15 recorded frames of each motility assay were conducted in the absence of any motility buffer (containing A T P ) . Following these control frames, the motility buffer containing A T P was added to the motility chamber via hanging drop and drawn through via a hanging piece of filter paper (Figure 8) in order to not change the focal plane. For microtubule movement on glass In order to verify that the movement recorded was in fact on the isolated junction plaque and not on the glass coverslip due to any motors from the spermatid/ectoplasmic specialization preparation or the additional testicular cytosol, one picture was taken in 68 the plane of the coverslip to see if there was in fact a separate and distinct population of microtubules present other than those found attached to the ectoplasmic specialization. For temperature The majority of experiments were conducted at room temperature (about 20°C-25°C) , but a limited number were conducted at 33 °C (testicular temperature) to see if there was any difference in the rate of microtubule movement. The temperature increase was obtained through the use of a stage heater which consisted of a 2mm aluminum plate (7.5 X 4.5 cm) with a central circular 2cm in diameter hole in it (Beach and Vogl , 1999). The base of this heater consisted of a circular heating element along with a thermistor probe (both purchased from Wellesely Hills, M A . ) to detect the stage temperature (Beach and Vogl , 1999). The glass motility chamber was added to the aluminum plate through the use of heat-seal grease (Beach and Vogl , 1999). The presence of cytosol in the motility buffer Experiments to see if cytosol could support motility of polar microtubules on the glass slide To determine if testicular cytosol in the motility buffer supported movement on kinesin and to determine if the testicular cytosol itself could support microtubule movement on glass, kinesin (Cytoskeleton, Denver, CO) or testicular cytosol (which presumably contained both positively and negatively directed motors) was added to the motility chamber at 5mg/ml for a 10 minute incubation at room temperature. Following this, 69 blocking buffer (containing 2 mg/ml casein) was added to the chamber for 10 minutes in order to block any non specific sites on the glass. In addition to this, experiments were run with both the inclusion and exclusion of 2 mg/ml testicular cytosol present in the motility buffer to see if there was any effect of the rate on microtubule movement. Experiment to see if microtubule motility occurred in the absence of any exogenous motor adhered to the motility chamber, but with testicular cytosol in the motility buffer. To determine if there was any movement due to the testicular cytosol in the motility buffer consisted of the exact same motility assay protocol as was used with the isolated ectoplasmic specializations, but was done without any of the cells. To a 10 pi aliquot of polar microtubules, 5 pi of PEM/250 was added and spun on a rotator for 10 minutes. Once completed, 10 pi of this solution was added to a motility chamber for another 10 minute incubation after which 10 pi of warm wash buffer was added. As was done in the motility assays using isolated ectoplasmic specializations on spermatids, the motility assay was run and motility buffer (containing 2mg/ml testicular cytosol) was added to the motility chamber after about 15 frames had elapsed. 70 Figure 7. Polarity marked G M P C P P microtubules bound to the ectoplasmic specialization. A phase image followed by images of actin and endoplasmic reticulum (to show the presence of the ectoplasmic specialization) and the G M P C P P polarity marked microtubules attached to the junction plaque as well as an overlay of the Polar microtubules on the ectoplasmic specialization. Actin = Red, Endoplasmic Reticulum = Blue, Polar Microtubules = Green. 71 Pha l l ac id i n Figure 8. Experimental method. Once focussed on the polar microtubule either attached to the glass for the control runs or the cell for the experimental runs, 10 pi of motility buffer (or stain) is added to the motility chamber via a hanging drop. The liquid is then removed via a hanging piece of filter paper by capillary action to not lose the focal plane of the polarity marked microtubules. Diagram was modified from Beach and Vogl (1999). 73 1H Data Analysis A l l still images were manipulated, without altering the integrity of the data, using Adobe Photoshop 5 (Adobe Systems Incorporated, San Jose, CA) . Rates A l l rates were calculated as done previously by Beach and Vogl (1999). The rates were determined by the distance that a polar microtubules moved in a continuous manner over the time that it took to process the frames of recorded movement. The rates are shown as a mean (um/sec) +/- standard deviation. Statistics Statistical tests for variance consisted of a two-tailed T-Test assuming unequal variances. Results from this test are routinely considered significant if their p-value is <.05. A p-value <. 1 is commonly considered a trend. 75 Immunofluorescent Experiments for Kinesin Preparation of Tissue Sections Fixation for sectioned material A l l animals were anesthetized using halothane and euthenized under deep anesthesia. Testes were removed from anesthetized animals and the spermatic artery was then canulated using a 26-gauge needle attached to a perfusion apparatus. The testes were perfused with phosphate buffered saline (PBS) at 33°C for 1-2 minutes to clear the organ of blood and was then perfused with 3.0% paraformaldehyde in PBS at 33°C. Sectioning Once the testis was fixed, it was then frozen in O C T compound and sectioned into 5 pm-10pm sections. The sections were then attached to polylysine coated slides and then plunged into cold acetone ( -20°C) with liquid nitrogen for 5 minutes to permeabilyze, dehydrate and further fix the tissue. The slides were then air-dried. This study was done 5 times and a total of 5 animals were used. 76 Preparation of isolated ectoplasmic specializations from tissue for kinesin immunofluorescence Tissue preparation of isolated ectoplasmic specializations for immunofluorescent studies was accomplished in the same manner as the ectoplasmic specialization isolation for the motility assay. Differences in the protocols occur after the cells have passed through the sucrose gradients for the isolation of the late stage spermatids. Once the tissue has been spun through the sucrose gradients the layer containing the same stages of spermatids (along with their attached ectoplasmic specializations) were removed from the centrifuge tube by a 26 gauge needle and were re-suspended in 1ml of 3% paraformaldehyde for 10 minutes. Following this, the cells were spun twice in an Eppendorph desk-top centrifuge for 2 minutes at a low setting of 2. The tube containing the cells was rotated 180° between the two spins. After this, the cytosol was removed and the cells were gently re-suspended in 1ml of PBS where they stayed for another 10 minutes at which point they were also centrifuged two times for 2 minutes at a setting of 2 on the desk-top Eppendorph centrifuge. The cytosol was removed and 70pl of PBS was added to the pellet of cells and the cells were again gently re-suspended. Then 10 pi of cells were added to polylysine coated slides, the excess liquid was removed using filter paper and the cells were placed into cold acetone ( -20°C) for about 5 minutes. After this, the slides were air dried and placed in a humidity chamber. The cells were now ready for blocking solution. 77 Immunofluorescence of Kinesin Labeling for kinesin was accomplished as follows. A 5% normal goat serum (NGS) in T P B S / B S A (PBS containing .05% Tween-20 and .1% Bovine Serum Albumin) was added to the tissue and incubated at room temperature for 20 minutes to block against secondary antibody nonspecific binding. Following this, 50uL of a 1:50 dilution of a rabbit-anti-kinesin antibody in 1%NGS/TPBS/BSA was added as the primary antibody against kinesin (a polyclonal antibody to HIPYR, a peptide against the conserved heavy chain of kinesin and kinesin-like proteins generated from a synthetic peptide C H I P Y R E S K L T ) (BabCo, Berkley, California, U S A ) and was incubated overnight in a 4°C cold room. The slides were washed three times (10 minutes each wash) with T P B S / B S A . Then the secondary antibody was added to the tissue for a 1 hour incubation at 37°C. The secondary antibodies were a 1:100 dilution in T P B S / B S A of a goat anti-rabbit antibody conjugated either to FITC or Texas Red. Once completed, the tissue was again washed 3 times (10 minutes each), then mounted in Vectashield (pH 8.2). 78 Immunofluorescent Controls for Kinesin Controls included the following: to control for the specificity of the primary antibody, normal rabbit IgG (NRIgG) was made up to the same concentration as the primary antibody that was used in the experiment. The control for the secondary antibody and for auto-fluorescence both used 1% N G S / T P B S / B S A in place of the primary antibody. A l l slides used the same secondary antibody, except for the auto-fluorescence control, which used the T P B S / B S A buffer in its place. 79 Results Polar Microtubules In order to construct polarity-marked microtubules three methods were attempted (the "Cytoskeleton Kit" method, the " N E M " method and the " G M P C P P " method). The approach used for the polarity-marked microtubule motility assay was the G M P C P P method. Cytoskeleton Kit I was unable to construct polar microtubules using the kit from Cytoskeleton. The best product consisted of clumps of brightly and dimly labeled fluorescence (Figure 9). For the most part these clumps did not have a linear pattern and there were not any areas of apparent attachment of brightly and dimly labeled fluorescence (seeds to tails). When assayed for motility on a bed of kinesin, upon the addition of motility buffer, the clumps broke into fragments; although the fragments were motile, they were not suitable for use in my assay. 80 N E M Polar Microtubules The N E M polar microtubules initially did produce linear arrays of fluorescent microtubules that supported motility, but lacked seeds (Figure 10). Problems arose when the technique was optimized and polar microtubules were built. Initially the seeds depolymerized under the fluorescent beam (Figure 11). Once modified the seeds stuck to the glass surface (Figure 12) and following further modifications, they did support motility (in the absence of testicular cytosol) primarily (all but 1 polar microtubule) in the proper direction on a bed of kinesin at an average rate of .212 +/- .084 pm/sec (n=8) (Figure 13). The rate for the 1 minus-end directed moving polarity marked microtubule was .272 pm/sec (n=l). Regretfully, the yield of polar microtubules as opposed to non-polar microtubules (9:330) was too low for use in my motility assay. 81 Polar microtubules are in the appropriate direction on kinesin; and cytosol in the motility buffer increases rates G M P C P P Polar Microtubules The G M P C P P method consistently produced high quality polar microtubules. These polar microtubules had a clearly marked seed and had both a clearly marked long and short end (Figure 14). When run on a bed of kinesin, 35 of 37 polarity marked microtubules moved leading by their seed (Figure 15). The average rate of movement of the polar microtubules without the presence of testicular cytosol in the motility buffer was .097 +/- .033 urn/sec (n=20) for those leading by the seed and was .081 (n=l) for the single polar microtubule that moved trailing by the seed. The average rate of movement of the polar microtubules in the presence of the Beach batch of testicular cytosol in the motility buffer was .135 +/-.076 urn/sec (n=8) for those leading by the seed and was .211 (n=l) for the single polar microtubule that moved trailing by the seed. The T-test gave a p-value of .2090 when plus-end directed rates with no cytosol present in the motility buffer were compared to the plus-end directed rates of polar microtubule movement when the Beach batch of testicular cytosol was present in the motility buffer. This difference in rates was not significant. The average rate of movement of the polar microtubules with the presence of the Guttman batch of testicular cytosol in the motility buffer was .132 +/- .083 urn/sec (n=8) for those leading by the seed. There were none recorded that trailed by the seed. The T-82 test gave a p-value of .2893 when the rates with no cytosol present in the motility buffer were compared to the rates of polar microtubule movement when the Guttman batch of testicular cytosol was present in the motility buffer. This was again not significantly different. T-test results comparing motility rates of the Beach and Guttman batches of testicular cytosol in the motility buffers gave a p-value of .9290 which was also not a significant difference. 83 Cytosol supports movement in the plus-end direction The control runs on testicular cytosol adsorbed on to the motility chamber at 5mg/ml also supported motility and all of the polar microtubules recorded (n=158) led by their seed. The polar microtubules moved on the Beach batch of cytosol at an average rate of .455 +/- .118 pm/sec (n=33) when testicular cytosol was not present in the motility buffer (Figure 16). When the Beach batch of testicular cytosol was included in the motility buffer the average rate was .211 +/- .094um/sec (n=8). No polar microtubule movements trailed by their seed in any of the cytosol controls. The T-test gave a p-value of .00003 when the rates on the Beach testicular cytosol were compared with or without the presence of the Beach batch of testicular cytosol in the motility buffer (Figure 17). This was significantly different. The polar microtubules moved on the Guttman batch of cytosol at an average rate of .102 +/- .056 pm/sec (n=9) when testicular cytosol was not present in the motility buffer and when the Guttman batch of testicular cytosol was included in the motility buffer the average rate was .054 +/- .023 pm/sec (n=3). Again no polar microtubule movements trailed by their seed in any of the cytosol controls. The T-test gave a p-value of .0644 when the rates on the Guttman testicular cytosol were compared with or without the presence of the Guttman batch of testicular cytosol in the motility buffer (Figure 17). This was not a significant difference in rates but did show a trend. 84 Beach testicular cytosol controls were also performed at 33°C by using a stage warmer set at 37°C (to get a slide temperature of 33°C) . The average rate of polar microtubule movement without testicular cytosol present in the motility buffer was .140 +/- .084 um/sec (n=8). The rate when testicular cytosol was present in the motility buffer was .125 +/- .058 urn/sec (n=3). Again all of the polar microtubule movement led by the seed. The T-test gave a p-value of .7372 when the rates on the Beach testicular cytosol at 33 °C were compared with or without the presence of the Beach batch of testicular cytosol in the motility buffer. This was not a significant difference. When rates on the Beach batch of testicular cytosol were compared in the absence of testicular cytosol in the motility buffer at room temperature versus at 33°C the T-Test gave a p-value of .0000003 which is significantly different. This same experiment, now with testicular cytosol present in the motility buffer gave a p-value of .1151 which is not significantly different. No motility was detected in assays where polar microtubules were added to the motility chamber without previously adhering spermatid/junction complexes, kinesin, or testicular cytosol to the glass. Importantly, motility buffer in these experiments contained 2 mg/ml testicular cytosol. 85 Polar Microtubules Move in the Plus and Minus Directions on Isolated Ectoplasmic Specialization A total of 25 polar microtubule movements were recorded. O f these 25, 8 trailed by the seed and 17 led by the seed. Interestingly, the polar microtubule orientation was not in all cases in the same orientation relative to the spermatid head as in the Sertoli cell, in about half of the cases it was orientated in the opposite orientation to1 that found in Sertoli cells in vivo (Figure 18). In the majority of the situations a phase photograph was taken to see the orientation of the spermatid and actin and the endoplasmic reticulum were stained to see if they were present (Figure 7). In addition, photographs in the plane of the glass confirmed that the microtubules for which movements were recorded were on ectoplasmic specializations and not on the glass. Upon working out the motility protocol, about 1 motility event occurred in every 4 experiments. Since motility was immediately observed upon the inclusion of 2 mg/ml testicular cytosol to the motility buffer, the vast majority of motility assays were performed with its inclusion in order to standardize the experiments. 86 Movements Trailing by the Seed On isolated ectoplasmic specializations 8 out of the 25 recorded movements trailed by the seed (Figure 19). O f the eight movements, five rates could not be calculated due to technical problems in deciphering the exact frame that the movement either began or ended and/or the angle at which the polar microtubule moved along the cell. Two of the five movements occurred on isolated spermatids (with their ectoplasmic specializations) that were not spun through the sucrose gradients in order to concentrate the elongate spermatids. The three movements where the rates were calculated had an average rate of .085 +/- .031 um/sec (n=3). Movements Leading by the Seed On the isolated junction plaques 17 out of the 25 polar microtubule movements lead by the seed (Figure 18). These movements moved at an average rate of .062 +/-.066 um/sec (n=12) when cytosol was present in the motility buffer and at a rate of .105 um/sec (n=l) when testicular cytosol was absent from the motility buffer. Out of these 17, 4 movements could not be calculated due to technical difficulties. 87 Kinesin Immunofluorescence Immunofluorescence of the HIPYR kinesin antibody showed a staining pattern which localized around the head of the developing spermatid. On sectioned tissue, this fluorescence was brighter and seemed to be more orderly (along the acrosome area) from stages III to V with an apparent decrease in brightness by stage VII (Figure 20). On tissue where the spermatids along with their developing ectoplasmic specializations (similar to those used in the polar microtubule motility assay) were stained, similar patterns of fluorescence occurred (Figure 21). A western blot was not performed using this antibody on testis tissue. Controls Sectioned tissue (Figure 22 A-A' ) and isolated ectoplasmic specializations attached to the developing spermatids (Figure 23 A-A ' ) that were stained for kinesin only showed staining patterns similar to those seen previously (described above). Controls for the primary and secondary antibody specificity along with the autofluOrescent controls were all negative for both the sectioned tissue (Figure 22 B-D') and isolated ectoplasmic specializations (Figure 23-B-D'). 88 Figure 9 . Microtubules constructed using the Cytoskeleton Polar Microtubule Kit. Poorly polymerized dimly rhodamine labeled tubulin microtubules (#1 and #3) and brightly labeled rhodamine tubulin seeds (#2) lying on a bed of kinesin at room temperature. After the addition of A T P (frame #4) the microtubules broke apart into extremely small fragments of tubulin which did support motility but were not stable. 89 F i g u r e 10. N E M n o n - p o l a r i t y m a r k e d m i c r o t u b u l e s . L i n e a r ar rays o f N E M microtubules. Capab le o f mot i l i ty on a bed o f k ines in upon the addi t ion o f A T P at room temperature. A l t h o u g h capab le o f movemen t on a bed o f k i n e s i n , they were not suf f ic ient ly polar i ty labeled to use in m y assay system. 91 00:03 00:11 00:19 00:27 •' s ' x 3 2 v , . . 3 - " • . '''-- J -• , it-2 k • 1 ' ' 2 +ATP 8.5um 00:35 " • 00:43 00:51 00:59 i • $ 1 1 ' '2. . 1 ? -.2 01:07 01:15 01:23 01:31 1 i ? r. .... •• 1 2 01:39 i i 01:47 1 01:56 I 02:04 1 ; ' 2 "•; • 02:12 1 ' . ""2 • ' 2 02:20 2 02:28 2 • 02:36 Figure 11. N E M polarity marked microtubule seeds break apart. The N E M polarity marked microtubule brightly rhodamine tubulin labeled seeds depolymerize upon the addition of fluorescent light. The frames following the addition of A T P show N E M microtubules which are not polarity marked or motile on a bed of kinesin at room temperature. 93 00:11 , ^ 00:19 - P 00:27 - ' 00:35 » * 00:43 - ' 00:51 - * 00:59 '•"ff'.lt'iiHw . . , " 01:07 «• * 01:15 » * 01:23 » * 01:31 01:39 , * 01:47 • * 01:56 • * 02:04 02:12 i * 02:20 +ATP • * 02:28 'i 02:36 . - 02:52 - . * 03:08 . * 03:24 03:48 Figure 12. N E M polarity marked microtubule seed sticks to glass. Upon the addition of the Motility buffer containing ATP the brightly rhodamine tubulin labeled seed of the N E M polar microtubule sticks to the glass coverslip instead of moving across the glass on the bed of kinesin at room temperature. 95 0 0 : 0 3 5pm 0 2 : 5 2 0 3 : 4 0 0 4 : 2 0 \ 0 4 : 5 3 0 5 : 4 1 1 Figure 13. N E M polarity marked microtubules capable of motility. Motile N E M polar microtubules at room temperature on a bed of kinesin upon the addition of A T P . The majority of other motile microtubules are not polarity marked. 9 7 Figure 14. G M P C P P polarity labeled microtubules. A collage of various different polarity marked microtubules all containing a brightly rhodamine tubulin labeled seed and both a short, negative, dimly rhodamine labeled tubulin end and a long, positive, dimly labeled tubulin end. 99 too Figure 15. G M P C P P polar microtubule kinesin controls. Four separate fields of G M P C P P polarity marked microtubules on recombinant kinesin conducted at room temperature on the same slide. Polar microtubule movement leading by the seed is evident by following any of the numbered microtubules across their respective row. 101 1 v 00:03 \ * . . A T P S u m 1 00:19 V • * 1 00:35 1 Li 1 4 1 00:51 MS 3 00:56 4 ' I i)1:23 4 4 If # 01^9 4 • •>* - \ 01:56 4* 3 _ 02:44 7 * • 03.00 8 > • • 5 03:16 jp J * - £ i • > 7 - . • V5 V V 03:32 i s 5 6 ' 1 03:48 a . ,9 • 04:04 8 04:20 8 9 / *. — — 04:36 ' 9 8 V • \ 06:13 • \ \ j X ^ ° % 06:29 06:45 4 / * * v07:09 4 1 j v 1 0 , . Figure 16. G M P C P P polarity marked microtubules cytosol control at room temperature without the presence of testicular cytosol in the motility buffer. A representative sample of a two hour cytosol control experiment (Beach cytosol) in which all of the polarity marked microtubules that moved led by their seed. This motion can be tracked by following a numbered microtubule through the processive frames. 103 07:04 1 | / I + A T P _ 5 t m i _ 1 07:12 / 07:20 4 A 07:28 / 4 3 07:36 / 4 3 08:00 6 / 08:08 6 ^: y 08:16 V 6 7 08:24 7 08:33 5 / 7 08:41 8 / 08:49 8 08:57 9 8 09:05 / 8 9 09:13 9 09:21 / 8 9 09:29 / 8 09:37 / 8 — 9 09:45 f 10 9 09:53 / 10 10:01 / 10 10:10 ' / 10 10:18 < \ 10:26 10 Figure 17. Comparison of polar microtubule rates on the Beach and Guttman batches of testicular cytosol. A graphical representation of the average rates of polar microtubule motility on a bed of testicular cytosol in the absence and the presence of testicular cytosol in the motility buffer. Represents a significant difference; * Represents a trend. 105 101 Rate (|im/sec) CD B) 3 CD O cn -»• cn o to ro cn o CO cn o o cn m > O = o 3 co i | =* z CD O O o GO o O 2 o -j ° s = S ^ o O O CO O CO O CD -y o _ -< 0 3 -i L7> O > co Q O 1 r- o oo U CD CO 2. O I? = O 3 ' I 03 "0 5, S 3> co CD CD ^ 3 o c: o _ c 3 - d o o ^ CO O o cn O 0) O 5 C IT C_ CD 3 " CD H CD 3 T J CD 3 CD CD 0) CD O CD Q> 3 a > tr w CD 3 O CD H CD (A i f o" c O o (A O 0) #• 73 O O 3 Figure 18. Plus-end directed movement (small arrow shows the movement direction) on an ectoplasmic specialization upon the addition of A T P of a polarity marked microtubule (long end labeled +; short end labeled -) not in the same orientation as that found in vivo. The large arrow shows the seed of the polar microtubule of interest and the asterisks follows the seed throughout the movement sequence. Frames were exposed for three seconds each. Actin and D iOC6 (endoplasmic reticulum) were not recorded due to the cell's dissociation from the glass surface upon the addition of the two stains. 107 10? \ Figure 19. Minus-end directed movement on an ectoplasmic specialization (small arrow shows movement direction). Phase image of the spermatid followed by microtubule images where the microtubule movement trails by the seed (large arrow). The microtubule is oriented in the same orientation as that found in nature (long end is marked by +; short end is marked by -). Frames were exposed for three seconds each. Actin and D i O C 6 (endoplasmic reticulum) images were taken once the run was complete. 109 Phase f Actin r 00:11 2.5fim 00:43 00:59 01:31 03:15 +ATP 03:23 03:28 03:47 . + / ^ 03:55 04:03 04:11 04:19 y 04:27 " . / 04:35 04:43 04:51 04:59 / • ' 05:07 05:23 06:50 MTs on Glass > • //0 Figure 20. Paired micrographs of sectioned tissue showing the phase and the positive kinesin staining. The micrographs presented as a spermatogenesis stage progression. A - A ' : stage I-III; (A) A phase micrograph showing spermatocytes at the beginning of spermatogenesis. (A') Kinesin staining of the same tissue section showing kinesins concentration around an area consistent with the position of ectoplasmic specializations. B - B ' : stage IV-V; (B) a phase micrograph showing a later stage of spermatogenesis. Here the heads of the spermatids are beginning to concentrate and develop a slight curve. (B') Kinesin staining of the same tissue is present adjacent to the acrosome, with the majority of the staining present along the dorsal surface of the acrosome (this is where ectoplasmic specializations are known to occur). C - C : stage VI; (C) A phase micrograph showing a further stage in spermatogenesis. The developing spermatids are now further from the Sertoli cell nucleus and their heads have a greater curve. ( C ) Kinesin staining is present along the dorsal side of the acrosome of the developing spermatid, with a lesser amount of staining along the ventral side of the acrosome. D-D' : stage VII; (D) A phase micrograph showing a stage of spermatogenesis where the spermatids (large arrow) are at the apical region of the Sertoli cell. (D') Kinesin staining is virtually undetectable with these late spermatids but is present at sites consistent with the ectoplasmic specialization on the next population of developing spermatids (small arrows). E - E ' and F-F': stage IX-II; (E and F) A phase micrograph showing a stage of spermatogenesis where the spermatids are now elongate. (E' and F') Kinesin staining is present along both the dorsal and ventral surfaces of the early stage spermatid acrosomes. This area is consistent with the position of the ectoplasmic specialization. Figure 21. Micrographs of spermatids isolated in the same way as that used for the motility assay. Phase, actin, kinesin, and an overlay of the actin and kinesin are all showed as a spermatogenesis stage progression. A -A ' " : stage IV; (A) a phase micrograph of a spermatid at approximately stage IV. (A') actin staining of the same cell used to indicate the presence of the intact ectoplasmic specialization on this cell. (A") kinesin staining of the cell showing the presence of kinesin around the entire cell, with the majority of staining along the dorsal surface of the acrosome. (A'") color overlay of actin (red) and kinesin (green) showing the co-localization of both actin and kinesin around the acrosome of the developing spermatid (yellow). B-B'": stage V ; (B) a phase micrograph of a spermatid at stage V . (B1) actin staining of the same cell used to indicate the presence of the intact ectoplasmic specialization on this cell. (B") kinesin staining of the cell showing the presence of kinesin along the dorsal surface of the spermatid acrosome. (B'") color overlay of actin (red) and kinesin (green) showing the co-localization of both actin and kinesin along the dorsal surface of the acrosome of the developing spermatid (yellow). C - C " : stage VI; (C) a phase micrograph of a spermatid at stage VI. (C) actin staining of the same cell used to indicate the presence of the intact ectoplasmic specialization on this cell. (C") kinesin staining of the cell showing the presence of kinesin along the dorsal surface of the acrosome. (C") color overlay of actin (red) and kinesin (green) showing the co-localization of both actin and kinesin along the dorsal surface of the acrosome of the developing spermatid (yellow). 113 HH Figure 22. (A-D) Phase micrographs showing tissue that was used for the control data for the sectioned tissue. (A') Single labeled for kinesin micrograph showing the kinesin distribution expected. Note the labeling along the dorsal surface of the developing spermatid heads. (B') Kinesin negative control for primary antibody specificity. The primary rabbit anti-kinesin antibody was replaced by normal rabbit IgG (NRIgG) and resulted in no consistent pattern of labeling. ( C ) Kinesin negative control for secondary antibody specificity. Here, the primary antibody was replaced with 1% normal goat serum (NGS) in TPBS/BSA and there was not positive staining found associated with the tissue. (D') Kinesin negative control for auto-fluorescence. Here, the primary antibody was replaced with 1% NDS in TPBS/BSA and the secondary antibody was replaced with the TPBS/BSA buffer. Again there was no specific positive staining. 115 116 Figure 23. (A-D) Phase micrographs showing tissue that was used for the control data for the isolated spermatids along with their overlying ectoplasmic specializations. (A' ) Single labeled kinesin micrograph showing the kinesin distribution expected. Note the labeling along the developing spermatid heads. (B') Kinesin negative control for primary antibody specificity. The primary rabbit anti-kinesin antibody was replaced by normal rabbit IgG (NRIgG) and resulted in no consistent pattern of labeling. ( C ) Kinesin negative control for secondary antibody specificity. Here, the primary antibody was replaced with 1% normal goat serum (NGS) in T P B S / B S A and there was not positive staining found associated with the tissue. (D') Kinesin negative control for auto-fluorescence. Here, the primary antibody was replaced with 1% N D S in T P B S / B S A and the secondary antibody was replaced with the T P B S / B S A buffer. Again there was no positive results found. None of these sections were controlled for actin. 117 Discussion In this study I provide evidence that Sertoli cell junction plaques (ectoplasmic specializations) possess the ability to translocate polar microtubules in vitro in both the plus and minus directions. This is consistent with the microtubule-based spermatid translocation hypothesis. Moreover, evidence presented here is the first that supports the argument that a kinesin-like motor is associated with these junction plaques. These findings, in conjunction with the finding that dynein occurs in Sertoli cell regions containing the plaques (Miller et al., 1999) could account for the movement of spermatids during spermiogenesis in vivo. The microtubule-based spermatid translocation hypothesis states that spermatids are moved in the seminiferous epithelium via the transport, along microtubule tracts, of unique Sertoli cell junction plaques attached to spermatid heads. These Sertoli cell junction plaques are composed of (moving cytoplasmically) hexagonally packed actin bundles and an attached cistern of endoplasmic reticulum. Anchored to the cytoplasmic face of the endoplasmic reticulum are thought to be molecular motors that move along Sertoli cell microtubules (oriented parallel to the long axis of the Sertoli cell) and thereby move the developing spermatids, within a Sertoli cell crypt, towards the base of the seminiferous epithelium and back up to the apex. If this hypothesis is true, then when polarity-labeled microtubules are added to isolated junction plaques (ectoplasmic specializations) and assayed for motility, both plus-end and minus-end directed movements should be observed. This would account for the movement thought to occur in vivo. 119 Establishment of the assay to verify the prediction: In order to perform polarity-marked microtubule assays in any system certain factors must be overcome: 1) Microtubules used in the assay must be visibly polar (the seed must be brighter than the ends). 2) The seeds of the polar microtubules must not be too "sticky" (or else they would not move). To acquire evidence in support of the bi-directional movement of microtubules over ectoplasmic specializations a number of unique conditions had to be achieved in the assay system 1) The microtubules must be concentrated. 2) The microtubules must be an appropriate size to bind, but not break or be too long for the cell. 3) The polar microtubules must be bright enough to be recorded through a full neutral density filter 4 ) Al l of the components of the ectoplasmic specialization must be present on the isolated spermatids including the actin and the endoplasmic reticulum 5) The polar microtubules must bind to the junction plaque and not extend onto the glass. 120 6) Binding must "favor" molecular motors and not any linking or binding elements. 7) The motors can only be of one type per polar microtubule (either only plus-end or only minus-end directed motors). 8) The cell can not "float away" once the motility buffer is added. 9) Polar microtubules must move a minimum of .5pm in order to be able to be detected. 10) The plane of the polar microtubule of interest must stay in focus throughout the movement sequence and both ends of the microtubule should be visible at some point during the motility event. 121 Polarity-marked microtubules The N E M polar microtubules were the preferred polar microtubule type. This was due to their non-ambiguous nature. There was no question as to polarity since N E M is not able to polymerize tubulin onto the negative end of the microtubule. Although preferred, this method was not used for the following reasons: (1) The incredibly low polar microtubule to non-polar microtubule yield (9 polar to 330non-polar); (2) The time that it took to grow the polar microtubules (6 hours for the seeds; 1 hour for the tail); (3) The stickiness of the seeds; (4) The inconsistent polymerization results and; (5) The indication that N E M may interfere with the microtubule lattice structure. The polar microtubules finally used were the G M P C P P polar microtubules. These polar microtubules were (1) consistently polarity labeled, (2) of uniform length, (3) could be concentrated and (4) took about 5 minutes to construct. They were assayed on a bed of kinesin to verify the proper position of the seed. This was performed by observing the polarity-labeled microtubules moving in the appropriate direction (leading by the seed) on the bed of kinesin. Once the polar microtubule construction had been overcome, these microtubules had to attach to the isolated ectoplasmic specializations. By concentrating the polar microtubules in a small amount of the same buffer as that used throughout the entire assay (PEM/250), blocking non-specific sites on the isolated ectoplasmic specializations by re-suspending the concentrated spermatids with the overlying ectoplasmic 122 specializations in 5mg/ml blocking buffer, adding the ectoplasmic specializations to the polar microtubules and placing this mixture on a rotator (to increase the number of polar microtubules that could "hit" the ectoplasmic specializations) for 10 minutes, I was able to attach the G M P C P P polarity-marked microtubules to the isolated ectoplasmic specializations. Testicular cytosol was used in this assay for a number of reasons. (1) Previous molecular motor work indicated an increase in binding through the use of 2mg/ml of cytosol (Blocker et al., 1996). (2) Other work showed that motility was obtained only if cytosol was added into the motility system (Schroer et al., 1988; Muresan et al., 1996) and (3) my control experiments revealed that the presence of cytosol in the motility buffer did not abolish motility (although the rates tended to be slower). I used a crude testicular cytosol rather than isolating Sertoli cell cytosol because of the many problems associated with isolating a Sertoli cell only cytosol. 123 Ectoplasmic specializations attached to isolated spermatids translocate polar microtubules in both the plus and minus directions Through the use of this polar microtubule motility assay, evidence provided in this thesis indicates that motility does occur in both the plus and minus directions. O f the 25 motility events, 17 were in the plus direction and 8 were in the minus direction. The findings that spermatid junction complexes support microtubule transport confirms earlier results by Beach and Vogl (1999). Moreover, the 8 minus-end directed motility events are consistent with the immunofluorescent findings of Miller et al. (1999) of dynein at the ectoplasmic specializations through the use of an antibody raised against the 74 kD dynein intermediate chain. M y findings strongly support the argument that a kinesin-like motor protein also is associated with the plaque. A possible reason that the majority of recorded movements were in the plus-end direction is that kinesin directed movement may dominate when both kinesin and dynein are present on cargo. Muresan et al (1996) found that the direction of vesicle movement appeared to be regulated by the presence or absence of a tightly bound plus-end kinesin motor (Muresan et al., 1996). That is, vesicles moved towards the minus-end only when a minus-end directed motor was bound to the vesicles in the absence of a plus-end directed motor (Muresan et al., 1996). Kinesin appears to override cytoplasmic dynein when both motors are bound to beads (Muresan et al., 1996). Perhaps one of the most significant findings of my study is that I was able to demonstrate movement in both directions, although the majority of movements were in the kinesin (plus-end) direction. 124 Since cytosol adsorbed onto the slide did support motility, it was important to verify that the movement was on the ectoplasmic specializations and not on the glass. In order to confirm this (1) actin was stained for, (2) the endoplasmic reticulum was stained for and (3) a photo in the plane of the glass was also taken at the end of the run. In some cases (4 out of the 25 movements) upon the addition of the stains, the isolated spermatid was washed off of the slide and therefor images of the actin and endoplasmic reticulum were not taken. The rates on the isolated spermatids with their exposed ectoplasmic specializations were relatively slow compared to those generally accepted for purified kinesin or dynein and were about 2 times slower than those recorded on purified kinesin which were performed in this study. Reasons for this could be that (1) this is not a purified system, (2) linking or binding elements could be present thereby slowing down the motility rates, (3) controlling factors could still be present in the assay system, (4) components of the constructed polar microtubules could be interfering with the molecular motors or (5) the motors could be a new type of motor or a new isoform of kinesin or cytoplasmic dynein which move at slower rates. There were no significant differences found between the rates of the plus versus minus end directed movements. A reason for this could be the small sample size. 125 Temperature Temperature was not controlled. A l l ectoplasmic specialization motility experiments were conducted at room temperature instead of at testicular temperature (33°C) since motility rates of polar microtubules on the Beach batch of cytosol were significantly slower at 33°C than those at room temperature. Also, motility on testicular cytosol stops on elevation of temperature (Beach and Vogl , 1999). Interestingly, a recent paper by Vaughan et al, (1999) showed that staining patterns of the dynein receptor, dynactin, were increased by incubating their cells for 1 minute at room temperature before fixing them at 37°C. This result indicates that there may be an effect of temperature on the receptors for the motors. The issue of temperature effects on motility in this system has not been properly resolved, or even addressed yet. 126 Evidence that the observed microtubule transport occurs on ectoplasmic specializations and is specific The motility events that have been discussed thus far are consistent with the conclusion that molecular motors are present on the endoplasmic reticulum of the ectoplasmic specializations. Evidence for this comes from a variety of observations. First, upon the completion of the motility run, the isolated spermatids were stained with phallacidin and D iOC6 to visualize the actin bundles and the endoplasmic reticulum respectively. Even though the D i O C 6 dye is not specific for only the endoplasmic reticulum, its staining pattern is consistent with the location of the endoplasmic reticulum as seen by electron microscopy (Vogl, 1996). Second, all of the movements on the testicular cytosol were in the plus-end direction, where on the ectoplasmic specializations, the movement was present in both the plus and minus directions. Third, although experiments without the presence of testicular cytosol in the motility buffer were seldom conducted, I did record one movement under these conditions on the ectoplasmic specialization in the plus direction, which further strengthens the concept of the motion being due to motors present on the plaque. Beach and Vogl (1999) have previously demonstrated that movement on the ectoplasmic specialization of microtubules occurs in the absence of cytosol in the motility buffer. Fourth, any polar microtubules added either to the ectoplasmic specializations or the glass did not move in the absence of A T P . 127 The data presented here strongly support the presence of a plus-end directed kinesin-like motor on the ectoplasmic specialization. The evidence for dynein is somewhat less strong because the possibility of breakage cannot be completely eliminated. 128 Evidence from immunofluorescence indicates the presence of a kinesin-like motor at ectoplasmic specializations Through immunofluorescent staining of sectioned tissue I found evidence of a kinesin-like protein at Sertoli cell regions known to contain ectoplasmic specializations. This was further corroborated by the same staining pattern present at regions consistent with ectoplasmic specializations on tissue isolated in the same manner as that used in the polar-microtubule motility assay. Although a western blot was not performed using this antibody (due to the lack of antibody), these results together with the motility data strongly indicate the presence of a kinesin-like protein at ectoplasmic specializations which have a likely function of moving the developing spermatids from the apex of the Sertoli cell towards the Sertoli cell nucleus. 129 Summary The most significant finding in this study is that translocation of polar microtubules on isolated ectoplasmic specializations in vitro occurs both in the plus and in the minus direction. The data are consistent with the microtubule-based spermatid translocation hypothesis, which hypothesizes that molecular motors are attached to the endoplasmic reticulum of the ectoplasmic specialization and is also consistent with the direction of spermatid translocation in vivo. This motility data together with the preliminary kinesin immunofluorescent results and the previous dynein (74 kD intermediate chain) evidence from Miller et al (1999) strongly suggest that a kinesin is likely the motor involved in translocating the developing spermatids from the apex of the seminiferous epithelium towards the base and a cytoplasmic dynein is likely the motor involved in returning the spermatids to the apex for eventual release into the lumen of the seminiferous tubule. 130 C o n c l u s i o n s 1) A p o l a r i t y -ma r k e d m i c r o t u b u l e m o t i l i t y a s say has n o w bee n d e v i s e d f o r the s t u d y o f m o l e c u l a r mo t o r s o n e c t o p l a sm i c s p e c i a l i z a t i o n s i n v i tro. 2) E c t o p l a s m i c s pe c i a l i z a t i o n s are c a p ab l e o f t r a n s po r t i n g p o l a r m i c r o t u b u l e s i n b o t h the p l us-end a n d m i n u s-end d i r e c t i o n s i n an in v i t r o mo t i l i t y assay. 3 ) T h e m o t i l i t y d a t a a n d p r e l i m i n a r y i m m u n o f l u o r e s c e n t d a t a c o n s t i t u t e the first e v i d e n c e t ha t a k i n e s i n - t y p e m o t o r m a y b e a s s o c i a t e d w i t h e c t o p l a s m i c s pec i a l i z a t i o n s . 4) R e s u l t s are cons i s ten t w i t h the m i c r o t ub u l e-ba sed s p e rma t i d t r a n s l o c a t i o n h y p o t he s i s . 131 F u t u r e d i r e c t i o n s I n o r d e r t o u n d e r s t a n d t h e e n t i r e p r o c e s s o f s p e r m a t o g e n e s i s as i t r e l a t e s t o t h e m i c r o t u b u l e - b a s e d s p e r m a t i d t r a n s l o c a t i o n h y p o t h e s i s n u m e r o u s o t h e r e x p e r i m e n t s s h o u l d b e c o n d u c t e d . I n c l u d e d a m o n g s t t h e s e a r e (1) d r u g s t u d i e s ; b l o c k i n g t h e m o t i l e a b i l i t y o f e i t h e r k i n e s i n o r d y n e i n a n d s e e i n g i f a l l o f t h e m o t i l i t y o b s e r v e d i s i n t h e d i r e c t i o n o f t h e n o n - b l o c k e d m o t o r , (2) s t a g e d e p e n d e n t p o l a r i t y - m a r k e d m i c r o t u b u l e m o t i l i t y a s s a y s ; m o r e p l u s - e n d d i r e c t e d m o v e m e n t s h o u l d o c c u r f r o m s t a g e s I I I - V a n d m o r e m i n u s - e n d d i r e c t e d m o v e m e n t s h o u l d o c c u r f r o m s t a g e s V - V I I , (3) a n t i b o d i e s s h o u l d b e r a i s e d a g a i n s t t h e s e e c t o p l a s m i c s p e c i a l i z a t i o n s p e c i f i c k i n e s i n - l i k e a n d d y n e i n - l i k e m o t o r s , (4) t h e p l u s a n d m i n u s e n d d i r e c t e d m o t o r s s h o u l d b e s e q u e n c e d a n d (5) t h e i m m u n o e l e c t r o n m i c r o s c o p y o f t h e k i n e s i n - l i k e p r o t e i n s h o u l d b e d o n e . 132 References Aizawa, H., Sekine, Y . , Takemura, R., Zhang, Z. , Nangaku, M. , Hirokawa, N. 1992 Kinesin superfamily in murine central nervous system. 1992 J. 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