actin-based endocytic machinery associated with junction turnover in the seminiferous epithelium.

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Contents lists available at ScienceDirect

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Review

Novel clathrin/actin-based endocytic machinery associated with junction turnover in the seminiferous epithelium A. Wayne Vogl a,∗ , Min Du a , Xue Ying Wang a , J’Nelle S. Young b a b

Department of Cellular and Physiological Sciences, Faculty of Medicine, University of British Columbia, Vancouver, BC Canada V6T 1Z3 Department of Radiology, College of Human Medicine, Michigan State University, Grand Rapids, MI 49503, USA

a r t i c l e

i n f o

Article history: Available online xxx Keywords: Tubulobulbar complex Junction endocytosis Sperm release Spermatocyte translocation

a b s t r a c t Tubulobulbar complexes are elaborate clathrin/actin related structures that form at sites of intercellular attachment in the seminiferous epithelium of the mammalian testis. Here we summarize what is currently known about the morphology and molecular composition of these structures and review evidence that the structures internalize intercellular junctions both at apical sites of Sertoli cell attachment to spermatids, and at basal sites where Sertoli cells form the blood–testis barrier. We present updated models of the sperm release and spermatocyte translocation mechanisms that incorporate tubulobulbar complexes into their designs. Crown Copyright © 2013 Published by Elsevier Ltd. All rights reserved.

Contents 1.

2.

3.

4.

5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Spermatogenesis and organization of the seminiferous epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Organization of intercellular junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Junction remodeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tubulobulbar complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. General morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Molecular components . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Apical tubulobulbar complexes and sperm release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Arrangement and position of apical tubulobulbar complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Apical tubulobulbar complexes and junction internalization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Sperm release model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Basal tubulobulbar complexes and spermatocyte translocation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Arrangement and position of basal complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Basal tubulobulbar complexes and junction internalization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Spermatocyte translocation model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction 1.1. Spermatogenesis and organization of the seminiferous epithelium

∗ Corresponding author. Tel.: +604 822 2395; fax: +604 822 2316. E-mail addresses: [email protected] (A.W. Vogl), [email protected] (M. Du), [email protected] (X.Y. Wang), J’[email protected] (J.S. Young).

Spermatogenesis involves the differentiation of relatively generalized diploid cells (spermatogonia) into specialized haploid cells (spermatids) that are released as spermatozoa into the duct system of the male reproductive tract. The process occurs in one of the most complex epithelia of the body known as the seminiferous epithelium. This stratified epithelium consists of two populations of cells, Sertoli cells and spermatogenic cells. The Sertoli cells form

1084-9521/$ – see front matter. Crown Copyright © 2013 Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.semcdb.2013.11.002

Please cite this article in press as: Vogl AW, et al. Novel clathrin/actin-based endocytic machinery associated with junction turnover in the seminiferous epithelium. Semin Cell Dev Biol (2013), http://dx.doi.org/10.1016/j.semcdb.2013.11.002

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Fig. 1. Shown here is a diagram of the rat seminiferous epithelium illustrating the locations of tubulobulbar complexes. The figure progresses through spermatogenesis from stages IV/V on the left to stages VIII/IX on the right. Tubulobulbar complexes develop at apical sites where Sertoli cells are attached to spermatids and at basal sites where neighboring Sertoli cells are attached to each other. The appearance of tubulobulbar complexes is correlated with a reduction in intercellular junctions that ultimately results in sperm release at apical sites and spermatocyte translocation at basal sites.

the architectural elements of the epithelium. These cells are irregularly columnar in shape and extend from the base to the apex of the epithelium. Through an elaborate junction complex near their bases, they divide the epithelium into two compartments—a basal compartment below the junctions and an adluminal compartment above. The spermatogenic cells lie between and are attached to the Sertoli cells. These cells ultimately give rise to the male gametes during the process of spermatogenesis. The process begins with spermatogonia in the basal compartment. Through a series of mitotic divisions in the basal compartment, these cells give rise to spermatocytes that enter meiosis, cross the junction complexes between Sertoli cells and move into the adluminal compartment to complete meiosis and become haploid spermatids. These latter cells undergo a series of elaborate morphological changes to become mature cells that ultimately are released from the epithelium as spermatozoa. 1.2. Organization of intercellular junctions Morphologically identifiable intercellular junctions occur at two major locations in the epithelium and their function is absolutely essential to spermatogenesis (Fig. 1). The first location is near the base of the epithelium where massive homotypic junction

complexes consisting of adhesion, tight, gap and desmosome-like junctions link together neighboring Sertoli cells. The adhesion junctions are unique to the seminiferous epithelium and are termed ‘ectoplasmic specializations’. They are characterized by a layer of hexagonally packed actin filaments situated between the plasma membrane and a cistern of endoplasmic reticulum, and are mainly nectin- and integrin-based attachments [1–3]. Tight and gap junctions to some extent overlap with, or are contained within, regions occupied by ectoplasmic specializations. Desmosome-like junctions, that are predominantly intermediate filament related, are intercalated amongst the ectoplasmic specializations where they appear in discontinuities or breaks in the adhesion zones. Tight junctions within the basal junction complexes [4], together with the bodies of the Sertoli cells, form the ‘blood–testis’ or Sertoli cell’ barrier that sequesters post-meiotic spermatogenic cells in an immuno-privileged and physiologically distinct environment above the junctions. This adluminal compartment is essential for spermatogenic cells to progress through spermatogenesis. The second major location where junctions occur is between Sertoli cells and spermatids (Fig. 1). Here, the junctions are heterotypic and consist almost entirely of ectoplasmic specializations. As with ectoplasmic specializations in basal junction complexes, those at apical sites are nectin- and integrin-based [1–3,5]. Unlike




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at basal sites, the characteristic actin filaments and the cisternae of endoplasmic reticulum occur only on the Sertoli cell side of the junction. These junctions firmly anchor developing spermatids to Sertoli cells and participate in positioning spermatids in the epithelium. 1.3. Junction remodeling Major remodeling of intercellular junctions in the mammalian seminiferous epithelium is correlated with two events during spermatogenesis that are fundamental to male fertility: (1) the release of spermatids as spermatozoa from the epithelium, which involves the elimination of apical ectoplasmic specializations; (2) the translocation of spermatocytes from basal to adluminal compartments of the epithelium, which involves the movement of the cells through basal junction complexes (Fig. 1). Generally in other epithelia, the internalization of intercellular junctions involves the disengagement of junction molecules in the plasma membrane of one cell from those in the adjacent plasma membrane followed by the internalization of the junction proteins into each cell by conventional methods [6–8], often involving clathrin [9]. This also is true of cell/substrate attachments [10]. An exception to these more conventional mechanisms, where each cell internalizes its own junction proteins, occurs with gap junctions where one cell may internalize ‘intact’ junctions into large double membrane vesicles that contain the membranes and junction molecules from both cells [11–13]. Interestingly, this process also is clathrin mediated [14] even though the vesicle sizes are much larger than conventional clathrin associated pits and vesicles. The protein clathrin plays a major role in forming coated vesicles for endocytosis and intercellular trafficking of specific cargo generally in cells [15]. Clathrin is made up of three limbs that radiate from a central hub forming a triskelion. This unique conformation contributes to its membrane shaping ability when a number of individual clathrin proteins link together forming a dome-shaped lattice. Although clathrin does not bind directly to the membrane, adaptor proteins such as AP-2 bind clathrin to specific integral membrane proteins [16]. After forming a coated-pit, the forming clathrin-coated vesicle undergoes scission that separates it from the parent membrane and shortly after, the coat disassembles [17]. In the seminiferous epithelium, current evidence is consistent with the hypothesis that more conventional clathrin-based endocytosis machinery has evolved into elaborate subcellular machines termed ‘tubulobulbar complexes’ that internalize ‘intact’ intercellular junctions into Sertoli cells and that these subcellular machines play a significant role in junction remodeling. 2. Tubulobulbar complexes 2.1. General morphology Tubulobulbar complexes consist of elongate tubular extensions that project either from spermatids or from Sertoli cells into corresponding invaginations of adjacent Sertoli cells at sites of cell/cell junctions [18] (Fig. 1). The double plasma membrane cores are roughly 50 nm in diameter and the space between the two plasma membranes measures around 6–8 nm [18]. The membrane cores are cuffed by a network of actin filaments [19,20], and each complex is capped by a coated pit [20]. Significantly, the coated-pit remains on the end of the complex throughout the maturation of the complex and does not undergo scission. In addition, the coated-pit consists of the two plasma membranes of the attached cells and fine filamentous connections span the intercellular space between the two membranes of the pit. A large swelling or ‘bulb’

Fig. 2. Schematic diagram showing the parts and identified molecular components of fully developed tubulobulbar complexes. (ER-PM junction = endoplasmic reticulum-plasma membrane junction).

develops in the distal third of each tubulobulbar complex as it matures. The bulb lacks an actin cuff, but is closely associated with cisternae of endoplasmic reticulum [18]. Eventually the bulb buds from the complex forming a large double membrane vesicle that enters endocytic compartments of the Sertoli cell and is degraded [18,21]. 2.2. Molecular components Over the last few years, significant progress has been made in identifying molecular components of tubulobulbar complexes (Fig. 2). It is now know that clathrin [22], the adaptor protein AP-2 [23], and the AP-2 binding protein Eps15 [24] are constituents of the coated pits. Amphiphysin [25] and dynamin [25,26] occur along tubular regions of the complexes and are likely in direct association with the membrane cores. Generally in other systems, both of these




Fig. 3. Light and electron micrographs showing the location and structure of apical tubulobulbar complexes in the rat. (a) 1 ␮m thick plastic section of seminiferous epithelium at stage VII of spermatogenesis stained with toluidine blue. Tubulobulbar complexes appear as organized clusters of linear structures adjacent to the concave face of the hook-shaped spermatids at the apex of the epithelium. In favorable sections, two parallel rows of complexes can be observed adjacent to each spermatid head (b). (c) Electron micrograph showing tubulobulbar complexes (TBCs) in cytoplasm adjacent to a spermatid head. Parts of tubulobulbar complexes and related structures are shown at higher magnification in the electron micrographs shown in panels (d) and (e). Bars = (a) 10 ␮m, (b) 5 ␮m, (c) 500 nm, (d) 250 nm, top inset 100 nm, bottom inset 100 nm, (e) 500 nm.

proteins sense and produce membrane curvature [27,28], while dynamin also promotes fission of the necks of endocytic vesicles [28]. The actin cuff surrounding the membrane core includes the components characteristic of a dendritic actin network in general in addition to elements involved with regulating actin assembly or with linking one filament to another. Those that characterize dendritic actin assembly include Arp2/3 and N-WASP [22] while among other actin related proteins identified are cofilin [29], espin [21], paxillin [30], cortactin [22] and Eps8 [31]. Forming a shell around the actin cuff are elements of the spectrin/plectin cytoskeleton [32,33]. By analogy with other systems [34], this spectrin/plectin shell may function to support the actin

network, link one complex to another, or determine the spacing pattern of complexes when the complexes occur in clusters [32,33]. Little is known about the bulb region of tubulobulbar complexes except that it lacks the surrounding dendritic network of actin filaments and is closely associated with cisternae of endoplasmic reticulum. This close association of endoplasmic reticulum with the plasma membrane is reminiscent of endoplasmic reticulumplasma membrane junctions in other systems that function mainly in calcium signaling [35]. Although tubulobulbar complexes have a number of unique morphological features and molecular components, their basic structure and composition is remarkably similar to that described for the classic clathrin mediated endocytosis machinery in other




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Fig. 4. Model of sperm release. Sertoli cells anchor spermatids to the apex of the epithelium through large adhesion complexes known as ectoplasmic specializations. In preparation for spermatid detachment, tubulobulbar complexes form in regions where ectoplasmic specializations begin to disassemble. These complexes internalize into the Sertoli cell regions of the plasma membranes of both the spermatid and Sertoli cell containing intercellular adhesion molecules [these regions are colored brown (spermatid) and blue (Sertoli cell) in the figure]. The bulb regions of the complexes bud-off from the structures and enter endocytic compartments of the Sertoli cell.

systems (see [17,36]). In fact, representative components in most of the protein domains defined for mammalian clathrin mediated endocytosis [17,37] have now also been identified as components of tubulobulbar complexes.

3. Apical tubulobulbar complexes and sperm release

rats they are slightly longer at 2–3 ␮m [18] and in opossum they extend up to 6–8 ␮m [38]. In all species, the general architecture of mature apical tubulobulbar complexes is similar; that is, all have tubular and bulbar regions (Fig. 3c and d), coated pits cap the ends of the complexes (Fig. 3d) and the structures are associated with endocytic compartments in the Sertoli cell (Fig. 3e).

3.1. Arrangement and position of apical tubulobulbar complexes 3.2. Apical tubulobulbar complexes and junction internalization Apical tubulobulbar complexes are those that occur in association with maturing spermatid heads at the apex of the seminiferous epithelium (Figs. 1 and 3). The structures arise in areas where ectoplasmic specializations attach spermatid heads to Sertoli cells, and multiple generations of complexes develop adjacent to each spermatid head [20]. The position of tubulobulbar complexes varies between species based on the shape of spermatid heads [38]. In animals with hook-shaped spermatid heads, such as in rats and mice, the complexes form a cluster adjacent to the convex face of the hook (Fig. 3a). In rats, the structures occur in two parallel rows with as many as 24 complexes present at any one time [18] (Fig. 3b). In animals with spade shaped spermatid heads, such as in humans, tubulobulbar complexes are more widely distributed, but tend to concentrate around the rim of the each head [38,39]. Mature complexes in most animals are around 1–2 ␮m long [38], whereas in

Apical tubulobulbar complexes have long been suspected of internalizing the attachments between Sertoli cells and spermatids [40], and therefore of being a significant part of the sperm release mechanism. Apical tubulobulbar complexes develop only at sites of attachment between Sertoli cells and spermatids and appear as ectoplasmic specializations disassemble. Tubulobulbar complexes contain the integral membrane adhesion proteins nectin-2 (Sertoli cell) [21,41], nectin-3 (spermatid) [21,41] and ␣6␤1 integrin (Sertoli cell) [41,42] that also are present in the membranes at the pre-existing ectoplasmic specializations. Moreover, terminal parts of tubulobulbar complexes and associated vesicles that are positive for junction proteins also are positive for endosomal and lysosomal markers (EEA1, Rab5, LAMP1, SGP1) [21,41,42].




Fig. 5. Light and electron micrographs showing the location and structure of basal tubulobulbar complexes in the rat. (a) Shown here is a 1 ␮m thick plastic section of seminiferous epithelium at stage V of spermatogenesis. The position of basal junction complexes between neighboring Sertoli cells are visible as long linear densities near the base of the epithelium. (b) Shown here is an electron micrograph of basal junction complexes between three Sertoli cells. Sections through a number of tubulobulbar complexes (TBCs) also are visible in the image. (c) Illustrated here is a bulb and the proximal tubule of a basal tubulobulbar complex. At a lower magnification, this tubulobulbar complex occurs within a pocket or fold in the junction complex between two adjacent Sertoli cells. Bars = (a) 10 ␮m, (b) 500 nm, (c) 250 nm, inset 500 nm.

If tubulobulbar complexes are significant components of the sperm release mechanism, then preventing the development of the complexes or interfering with their structure/function should prevent or delay sperm release. In rats treated with 17B-estradiol, apical tubulobulbar complexes do not form and this is correlated with spermiation failure or the inability to release sperm cells from the epithelium [43]. Similarly, in amphiphysin knockout mice [25], tubulobulbar complexes fail to develop and spermiation failure occurs. In rats, knocking down cortactin by intratesticular injection of siRNA duplexes results in dramatically shorter tubulobulbar complexes and in delayed or failed sperm release [44]. Thus the experimental data is consistent with a significant role of tubulobulbar complexes in sperm release.

3.3. Sperm release model The process of spermiation or sperm release has been described as consisting of a number of phases beginning with the movement of spermatids from deep in apical invaginations or crypts in Sertoli cells to the apex of the epithelium and ending with the actual detachment of morphologically mature cells from the epithelium [45,46]. Our discussion here focuses on that phase of spermiation involving the elimination of adhesion junctions between the Sertoli cells and spermatids. The structural and experimental data reviewed above lead us to propose the following model for elimination of apical adhesion junctions and the detachment of spermatids (Fig. 4).




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Fig. 6. Quantification of basal junction components at stages V and VII in rat seminiferous epithelium. The quantification presented here generally was done as described by Du and co-workers [23]. Cryosections of perfusion fixed testes were labeled for each of actin, claudin-11, nectin-2, connexin-43, and N-cadherin and evaluated by fluorescence microscopy. Images were digitized and an index of the amount of junction component present at each of stage V and stage VII of spermatogenesis was obtained by selecting an area around the periphery of each tubule that contained basal junctions and then determining the numbers of pixels above a predetermined threshold. For most junction components, 8 tubules for each stage from each of 8 animals were evaluated. For N-cadherin, 8 tubules from each of 6 animals were quantified. A t-test was used to determine if the numbers of pixels above threshold were different between stages V and VII. The numbers of pixels above threshold were significantly different between stages V and VII for actin, claudin-11 and nectin-2, but were not different for connexin-43 and N-cadherin.

Cytoplasmic components of ectoplasmic specializations are disassembled at specific locations around the spermatid head and simultaneously tubulobulbar complexes begin to form at these locations. Initiation of the complexes occurs by the formation of coated pits at the Sertoli cell surface that retain a connection to the spermatid plasma membrane, possibly through the action of adhesion proteins that remain engaged across the intercellular space. It also is possible that engagement persists between the two adjacent membranes via receptors and ligands that are currently unknown. The coated pits recruit cytoskeletal and other machinery common to clathrin mediated endocytic processes in general, but the coated pits fail to undergo scission and the necks dramatically increase in length thereby forming the characteristic appearance of tubulobulbar complexes. As the structures elongate, regions of the plasma membranes that remain attached by adhesion proteins are drawn into the tubulobulbar complexes. The formation of bulbs near the ends of the complexes dramatically increases the amount of membrane that can be incorporated into the complexes. The subsequent ‘budding’ of the bulbs results in the internalization of the junction components into the Sertoli cell. The formation of tubulobulbar complexes reduces the amount of adhesion junctions at the interface between the Sertoli cell and spermatid head resulting in the progressive separation

of the two cells. Remnants of adhesion junctions at the last remaining interface between Sertoli cells and spermatids, or the formation of a new attachment between the two cells, forms a small disengagement complex [46,47]. When this last focal attachment between the cells is disassembled, spermatids are released as spermatozoa into the duct system of the male reproductive tract. Adhesion proteins that are internalized either are degraded or, in the case of the Sertoli cell proteins, may be recycled [42]. Tubulobulbar complex mediated junction internalization at apical sites simultaneously accomplishes two processes: (1) removal of the massive intercellular attachment between Sertoli cells and spermatids ultimately resulting in sperm release; (2) removal of adhesion molecules from spermatid plasma membranes that are no longer needed by mature spermatozoa. A corollary of the latter function would be that tubulobulbar complexes participate in remodeling the plasma membrane of the spermatid head as part of the process of spermiation. In fact, we have argued elsewhere [39] that removal of elements in the plasma membrane of spermatids that are no longer needed after release may have been the selection pressure for the evolution of tubulobulbar complexes for junction remodeling generally by mammalian Sertoli cells.




Fig. 7. Model of basal junction remodeling during spermatocyte translocation from basal to adluminal compartments of the seminiferous epithelium. Tubulobulbar complexes begin to internalize intercellular junctions well before the movement of spermatocytes off of the basal lamina and through basal junction complexes. The activity of tubulobulbar complexes reduces the amount of intercellular junction present above spermatocytes prior to beginning their translocation in stage VIII. New junctions begin to form below the spermatocytes during stages IX–X as these cells move adluminally.

4. Basal tubulobulbar complexes and spermatocyte translocation 4.1. Arrangement and position of basal complexes Basal tubulobulbar complexes occur in association with Sertoli cell junctions that separate basal from adluminal compartments of the epithelium (Figs. 1 and 5). They often occur in folds or pockets within the junction belts circling basal regions of the Sertoli cells [23] (Fig. 5b and c). Basal complexes have the same characteristic features as apical ones [18,39]; however, the coated pits at the end of the basal complexes lack membrane-associated densities unlike the coated pits of apical complexes that have distinct electron dense material underlying the spermatid plasma membrane portion of the pit. Although basal tubulobulbar complexes have been far less studied than apical ones, they are well described in the rat [18,23,48] and human [39] and are likely present in all mammals. 4.2. Basal tubulobulbar complexes and junction internalization A growing body of evidence indicates that basal tubulobulbar complexes internalize intercellular junctions in a similar

fashion to that at apical sites, and that junction proteins in adjacent plasma membranes remain engaged across the intercellular space as they are drawn into the internalization machinery. At the ultrastructural level, tight and gap junctions are visible in the structures [23,48] and immunofluorescence staining with probes for claudin-11, connexin-43 and nectin-2 reveal rod-like profiles associated with basal junction complexes that are consistent with the predicted appearance of tubulobulbar complexes [23]. Moreover, and like at apical sites, endocytic markers occur in regions of the epithelium containing tubulobulbar complexes [23]. Recently, tubulobulbar complexes have been identified in morphologically differentiated primary cultures of Sertoli cells. In this culture system, as in the animal, the structures occur in association with basal junction complexes, and contain junction proteins and morphologically identifiable intercellular junctions [23]. A key finding reported by Russell [48] is that basal tubulobulbar complexes appear most numerous in early stages of spermatogenesis (stages III to VI) well before spermatocytes begin to move upwards in the epithelium at stage VIII. If basal tubulobulbar complexes internalize and degrade intercellular junctions, then there should be more basal junction present at stage V (when tubulobulbar complexes are most apparent) than at stage VII (when




tubulobulbar complexes are degrading). This appears to be the case when tubules are probed for actin and claudin-11 as markers for ectoplasmic specializations and tight junctions, respectively [23], and thresholding techniques are used to evaluate the data. The results reported in our earlier work have been repeated with consistent results (Fig. 6); that is, the numbers of positive pixels remaining above a pre-determined threshold at basal junctions both for actin and for claudin-11 are significantly different between stage V and stage VII. The same is true for nectin-2 when used as a marker for ectoplasmic specializations. However, when connexin-43, a gap junction protein, and N-cadherin, a canonical adherens junction protein, are used as markers the difference between the two stages in the numbers of pixels remaining above threshold does not reach significance. Because we know that gap junctions actually do occur within tubulobulbar complexes, we suspect that the lack of significance in connexin-43 staining between the two stages may be due to the fact that we did not differentiate between connexin-43 at gap junctions between neighboring Sertoli cells and those junctions between Sertoli cells and early spermatogenic cells. The latter would not be expected to be internalized by tubulobulbar complexes and likely have a different dynamic of turnover. The results for N-cadherin, may be due to the same issue, or the protein could possibly be internalized by another mechanism. This issue remains to be clarified.

4.3. Spermatocyte translocation model The translocation of primary (leptotene) spermatocytes into the adluminal compartment serves to re-populate the epithelium with the next generation of germ cells that will complete meiosis and morphologically differentiate into sperm cells. In the rat, primary spermatocytes begin to migrate in late stage VIII of spermatogenesis and complete the process in stage XI. This migration has been beautifully described in a classic paper by Russell [49]. In this early iteration of the model that remains the accepted model for basal junction turnover in the mammalian seminiferous epithelium [50], junctions disassemble above leptotene spermatocytes as these cells move upwards in the epithelium while simultaneously new junctions assemble below—a mechanism that not only facilitates the translocation of spermatocytes, but also retains the immunological and physiological barrier between basal and adluminal compartments. The presence of junction proteins in tubulobulbar complexes, and the correlation between the peak appearance of tubulobulbar complexes and an apparent reduction in the amount of basal junction material present at basal sites has suggested to us the following updated model for junction remodeling associated with spermatocyte translocation [23] (Fig. 7). The remodeling of basal junction complexes begins well before the actual apical movement of spermatocytes. In rats, basal junctions are very large and form broad belts around the circumference of the Sertoli cells at stages IV and V. Tubulobulbar complexes dramatically increase in number at this time and begin the process of junction remodeling in anticipation of spermatocyte translocation at later stages of spermatogenesis. From stages V to VII junctions are removed and degraded by tubulobulbar complexes so that by the time spermatocytes lift off the basal lamina and begin to translocate apically in stage VIII, the amount of junction material present above the cells is dramatically reduced. As the spermatocytes move upwards, junctions remaining above the cells continue to be internalized, while at the same time new junctions form below. This model is attractive because it incorporates tubulobulbar complexes into the junction remodeling mechanism, accounts for the quantitative data presented by Russell [48] on the changes in tubulobulbar complex numbers during spermatogenesis, unifies the function of

9

tubulobulbar complexes with that at apical sites, and is consistent with the morphology.

5. Conclusions Current morphological, molecular and experimental evidence is consistent with the conclusion that tubulobulbar complexes are clathrin-based structures responsible for internalizing intercellular junctions in the seminiferous epithelium as part of the mechanism both of sperm release from the apex of the epithelium and of spermatocyte translocation through basal junctions between Sertoli cells. The junction internalization hypothesis accounts for the presence of tubulobulbar complexes at apical and basal sites in the epithelium, the presence of junctions and junction proteins within the structures, the association of early endocytosis markers with tubulobulbar complexes and their related vesicles, and the similarity of basic structure and composition of tubulobulbar complexes with clathrin-based endocytosis machinery present generally in cells. Although we have learned a great deal about tubulobulbar complexes in the last few years, there are a number of important issues that remain to be resolved and a number of new issues have arisen. Some of these are summarized as follows:

(1) The process of vesiculation and budding of tubulobulbar complexes is not well understood. Although it is known that the bulb regions separate or ‘bud’ from the complexes and enter endocytic compartments of Sertoli cells, it is not entirely clear what happens to tubular parts of the complexes, nor is it clear what happens to the coated-pit at the end of each complex as budding of the bulbs occurs, or what triggers the budding. (2) Nothing is known about the role of the endoplasmic reticulum at tubulobulbar complexes. The presence of cisternae of endoplasmic reticulum both as characteristic components of ectoplasmic specializations and as constant features of the bulb regions of tubulobulbar complexes indicates to us that these domains of the endoplasmic reticulum in Sertoli cells may play key roles in the function and regulation of junction remodeling in the seminiferous epithelium. (3) The detailed relationship between markers of early endocytosis and mature tubulobulbar complexes and their related vesicles has yet to be determined at the ultrastructural level. (4) If tubulobulbar complexes are involved with internalizing membrane from the heads of spermatids during sperm release, as appears to be the case, then the plasma membranes of spermatid heads are likely dramatically remodeled during the release phase of spermatogenesis. This has fairly profound implications for how plasma membrane domains of spermatozoon heads are established. (5) The role of tubulobulbar complexes in the turnover of Ncadherin needs to be clarified. (6) Although the link between tubulobulbar complexes and sperm release has been reasonably well documented, the link between basal tubulobulbar complexes and spermatocyte translocation needs to be more firmly established experimentally. (7) The way in which disassembly of ectoplasmic specializations and the assembly of tubulobulbar complexes is coordinated during spermatogenesis has yet to be determined.

Acknowledgments This work was funded by an NSERC Discovery Grant (no. 155397-13) to AWV.




References [1] Palombi F, Salanova M, Tarone G, Farini D, Stefanini M. Distribution of beta 1 integrin subunit in rat seminiferous epithelium. Biol Reprod 1992;47:1173–82. [2] Salanova M, Stefanini M, De Curtis I, Palombi F. Integrin receptor alpha 6 beta 1 is localized at specific sites of cell-to-cell contact in rat seminiferous epithelium. Biol Reprod 1995;52:79–87. [3] Ozaki-Kuroda K, Nakanishi H, Ohta H, Tanaka H, Kurihara H, Mueller S, et al. Nectin couples cell-cell adhesion and the actin scaffold at heterotypic testicular junctions. Curr Biol 2002;12:1145–50. [4] Dym M, Fawcett DW. The blood–testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium. Biol Reprod 1970;3:308–26. [5] Inagaki M, Irie K, Ishizaki H, Tanaka-Okamoto M, Miyoshi J, Takai Y. Role of cell adhesion molecule nectin-3 in spermatid development. Genes Cells 2006;11:1125–32. [6] Le TL, Yap AS, Stow JL. Recycling of E-cadherin: a potential mechanism for regulating cadherin dynamics. J Cell Biol 1999;146:219–32. [7] Marchiando AM, Shen L, Graham WV, Weber CR, Schwarz BT, Austin 2nd JR, et al. Caveolin-1-dependent occludin endocytosis is required for TNF-induced tight junction regulation in vivo. J Cell Biol 2010;189:111–26. [8] Shen L, Turner JR. Actin depolymerization disrupts tight junctions via caveolaemediated endocytosis. Mol Biol Cell 2005;16:3919–36. [9] Ivanov AI, Nusrat A, Parkos CA. Endocytosis of epithelial apical junctional proteins by a clathrin-mediated pathway into a unique storage compartment. Mol Biol Cell 2004;15:176–88. [10] Ezratty EJ, Bertaux C, Marcantonio EE, Gundersen GG. Clathrin mediates integrin endocytosis for focal adhesion disassembly in migrating cells. J Cell Biol 2009;187:733–47. [11] Berthoud VM, Minogue PJ, Laing JG, Beyer EC. Pathways for degradation of connexins and gap junctions. Cardiovasc Res 2004;62:256–67. [12] Gaietta G, Deerinck TJ, Adams SR, Bouwer J, Tour O, Laird DW, et al. Multicolor and electron microscopic imaging of connexin trafficking. Science 2002;296:503–7. [13] Larsen WJ, Tung HN, Murray SA, Swenson CA. Evidence for the participation of actin microfilaments and bristle coats in the internalization of gap junction membrane. J Cell Biol 1979;83:576–87. [14] Piehl M, Lehmann C, Gumpert A, Denizot JP, Segretain D, Falk MM. Internalization of large double-membrane intercellular vesicles by a clathrin-dependent endocytic process. Mol Biol Cell 2007;18:337–47. [15] Doherty GJ, McMahon HT. Mechanisms of endocytosis. Annu Rev Biochem 2009;78:857–902. [16] Kirchhausen T. Clathrin. Annu Rev Biochem 2000;69:699–727. [17] McMahon HT, Boucrot E. Molecular mechanism and physiological functions of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 2011;12:517–33. [18] Russell L, Clermont Y. Anchoring device between Sertoli cells and late spermatids in rat seminiferous tubules. Anat Rec 1976;185:259–78. [19] Vogl AW, Soucy LJ, Lew GJ. Distribution of actin in isolated seminiferous epithelia and denuded tubule walls of the rat. Anat Rec 1985;213:63–71. [20] Russell LD. Further observations on tubulobulbar complexes formed by late spermatids and Sertoli cells in the rat testis. Anat Rec 1979;194:213–32. [21] Guttman JA, Takai Y, Vogl AW. Evidence that tubulobulbar complexes in the seminiferous epithelium are involved with internalization of adhesion junctions. Biol Reprod 2004;71:548–59. [22] Young JS, Guttman JA, Vaid KS, Shahinian H, Vogl AW. Cortactin (CTTN), NWASP (WASL), and clathrin (CLTC) are present at podosome-like tubulobulbar complexes in the rat testis. Biol Reprod 2009;80:153–61. [23] Du M, Young J, De Asis M, Cipollone J, Roskelley C, Takai Y, et al. A novel subcellular machine contributes to basal junction remodeling in the seminiferous epithelium. Biol Reprod 2013;88:60. [24] Nicholls PK, Harrison CA, Walton KL, McLachlan RI, O’Donnell L, Stanton PG. Hormonal regulation of sertoli cell micro-RNAs at spermiation. Endocrinology 2011;152:1670–83. [25] Kusumi N, Watanabe M, Yamada H, Li SA, Kashiwakura Y, Matsukawa T, et al. Implication of amphiphysin 1 and dynamin 2 in tubulobulbar complex formation and spermatid release. Cell Struct Funct 2007;32:101–13. [26] Vaid KS, Guttman JA, Babyak N, Deng W, McNiven MA, Mochizuki N, et al. The role of dynamin 3 in the testis. J Cell Physiol 2007;210:644–54.

[27] Peter BJ, Kent HM, Mills IG, Vallis Y, Butler PJ, Evans PR, et al. BAR domains as sensors of membrane curvature: the amphiphysin BAR structure. Science 2004;303:495–9. [28] Ferguson SM, De Camilli P. Dynamin, a membrane-remodelling GTPase. Nat Rev Mol Cell Biol 2012;13:75–88. [29] Guttman JA, Obinata T, Shima J, Griswold M, Vogl AW. Non-muscle cofilin is a component of tubulobulbar complexes in the testis. Biol Reprod 2004;70:805–12. [30] Mulholland DJ, Dedhar S, Vogl AW. Rat seminiferous epithelium contains a unique junction (ectoplasmic specialization) with signaling properties both of cell/cell and cell/matrix junctions. Biol Reprod 2001;64:396–407. [31] Lie PP, Mruk DD, Lee WM, Cheng CY. Epidermal growth factor receptor pathway substrate 8 (Eps8) is a novel regulator of cell adhesion and the blood–testis barrier integrity in the seminiferous epithelium. FASEB J 2009;23:2555–67. [32] Upadhyay R, D’Souza R, Sonawane S, Gaonkar R, Pathak S, Jhadav A, et al. Altered phosphorylation and distribution status of vimentin in rat seminiferous epithelium following 17beta-estradiol treatment. Histochem Cell Biol 2011;136:543–55. [33] de Asis MA, Pires M, Lyon K. A network of spectrin and plectin surrounds the actin cuffs of apical tubulobulbar complexes in the rat. Spermatogenesis 2013;3:e25733. [34] Gad A, Lach S, Crimaldi L, Gimona M. Plectin deposition at podosome rings requires myosin contractility. Cell Motil Cytoskeleton 2008;65:614–25. [35] Carrasco S, Meyer T. STIM proteins and the endoplasmic reticulum-plasma membrane junctions. Annu Rev Biochem 2011;80:973–1000. [36] Conibear E. Converging views of endocytosis in yeast and mammals. Curr Opin Cell Biol 2010;22:513–8. [37] Taylor MJ, Perrais D, Merrifield CJ. A high precision survey of the molecular dynamics of mammalian clathrin-mediated endocytosis. PLoS Biol 2011;9:e1000604. [38] Russell LD, Malone JP. A study of Sertoli-spermatid tubulobulbar complexes in selected mammals. Tissue Cell 1980;12:263–85. [39] Vogl AW, Young JS, Du M. New insights into roles of tubulobulbar complexes in sperm release and turnover of blood–testis barrier. Int Rev Cell Mol Biol 2013;303:319–55. [40] Russell LD, Goh JC, Rashed RM, Vogl AW. The consequences of actin disruption at Sertoli ectoplasmic specialization sites facing spermatids after in vivo exposure of rat testis to cytochalasin D. Biol Reprod 1988;39:105–18. [41] Young JS, Guttman JA, Vaid KS, Vogl AW. Tubulobulbar complexes are intercellular podosome-like structures that internalize intact intercellular junctions during epithelial remodeling events in the rat testis. Biol Reprod 2009;80:162–74. [42] Young JS, Takai Y, Kojic KL, Vogl AW. Internalization of adhesion junction proteins and their association with recycling endosome marker proteins in rat seminiferous epithelium. Reproduction 2012;143:347–57. [43] D’Souza R, Pathak S, Upadhyay R, Gaonkar R, D’Souza S, Sonawane S, et al. Disruption of tubulobulbar complex by high intratesticular estrogens leading to failed spermiation. Endocrinology 2009;150:1861–9. [44] Young JS, de Asis M, Guttman JA, Vogl AW. Cortactin depletion results in short tubulobulbar complexes and spermiation failure in rat testis. Biol Open 2012;1:1–9. [45] Russell LD. Spermiation—the sperm release process: ultrastructural observations and unresolved problems. In: Van Blerkom J, Motta PM, editors. Ultrastructure of Reproduction. Boston: Martinus Nijhoff; 1984. p. 46–66. [46] O’Donnell L, Nicholls PK, O’Bryan MK, McLachlan RI, Stanton PG, Spermiation. The process of sperm release. Spermatogenesis 2011;1:14–35. [47] Beardsley A, Robertson DM, O’Donnell L. A complex containing alpha6beta1integrin and phosphorylated focal adhesion kinase between Sertoli cells and elongated spermatids during spermatid release from the seminiferous epithelium. J Endocrinol 2006;190:759–70. [48] Russell LD. Observations on the inter-relationships of Sertoli cells at the level of the blood–testis barrier: evidence for formation and resorption of Sertoli–Sertoli tubulobulbar complexes during the spermatogenic cycle of the rat. Am J Anat 1979;155:259–79. [49] Russell L. Movement of spermatocytes from the basal to the adluminal compartment of the rat testis. Am J Anat 1977;148:313–28. [50] Smith BE, Braun RE. Germ cell migration across Sertoli cell tight junctions. Science 2012;338:798–802.


Seminiferous Epithelium Cycle in Bombina orientalis.

Visualizing the functional architecture of the endocytic machinery.

Study of the seminiferous epithelium cycle in Eulemur hybrids.

Intercellular junction development in maturing rat seminiferous tubules.

Duration of the cycle of the seminiferous epithelium in the prairie vole (Microtus ochrogaster ochrogaster).

An Endocytic Scaffolding Protein together with Synapsin Regulates Synaptic Vesicle Clustering in the Drosophila Neuromuscular Junction.

Integrin-beta3 clusters recruit clathrin-mediated endocytic machinery in the absence of traction force.

Retinoid acid: the trigger for the cycle of the seminiferous epithelium in the adult testis?

Role of spermatogonia in the stage-synchronization of the seminiferous epithelium in vitamin-A-deficient rats.

CEACAM2-L on spermatids interacts with poliovirus receptor on Sertoli cells in mouse seminiferous epithelium.

Regulation of microtubule (MT)-based cytoskeleton in the seminiferous epithelium during spermatogenesis.

Ultrastructure of the seminiferous epithelium and intertubular tissue of the human testis.

Expression of the c-Kit receptor in germ cells of the seminiferous epithelium in rats with hormonal imbalance.

Stage-specific cellular regulation of inhibin alpha-subunit mRNA expression in the rat seminiferous epithelium.

Prolonged suppression of spermatogenesis by oestrogen does not preserve the seminiferous epithelium in procarbazine-treated rats.

Synchronization of the seminiferous epithelium after vitamin A replacement in vitamin A-deficient mice.

A-single spermatogonia heterogeneity and cell cycles synchronize with rat seminiferous epithelium stages VIII-IX.

The changes of stage distribution of seminiferous epithelium cycle and its correlations with Leydig cell stereological parameters in aging men.

Protective Effects of Antioxidants on Sperm Parameters and Seminiferous Tubules Epithelium in High Fat-fed Rats.

Testicular toxicity of methylmercury: analysis of cellular distribution pattern at different stages of the seminiferous epithelium.

Stage- and cell-specific gene expression and hormone regulation of the seminiferous epithelium.

Connexin 43 expression in Sprague-Dawley rat seminiferous epithelium after in utero exposure to flutamide.

Relative volume of Sertoli cells in monkey seminiferous epithelium: a stereological analysis.

Phosphoinositide turnover associated with synaptic transmission.