BioMed Research International

BioMed Research International / 2014 / Article

Review Article | Open Access

Volume 2014 |Article ID 635979 |

Giovanna Berruti, Chiara Paiardi, "The Dynamic of the Apical Ectoplasmic Specialization between Spermatids and Sertoli Cells: The Case of the Small GTPase Rap1", BioMed Research International, vol. 2014, Article ID 635979, 9 pages, 2014.

The Dynamic of the Apical Ectoplasmic Specialization between Spermatids and Sertoli Cells: The Case of the Small GTPase Rap1

Academic Editor: Nicola Bernabò
Received12 Dec 2013
Accepted19 Jan 2014
Published27 Feb 2014


Despite advances in assisted reproductive technologies, infertility remains a consistent health problem worldwide. Spermiation is the process through which mature spermatids detach from the supporting Sertoli cells and are released into the tubule lumen. Spermiation failure leads to lack of mature spermatozoa and, if not occasional, could result into azoospermia, major cause of male infertility in human population. Spermatids are led through their differentiation into spermatozoa by the apical ectoplasmic specialization (aES), a testis-specific, actin-based anchoring junction restricted to the Sertoli-spermatid interface. The aES helps spermatid movement across the seminiferous epithelium, promotes spermatid positioning, and prevents the release of immature spermatozoa. To accomplish its functions, aES needs to undergo tightly and timely regulated restructuring. Even if components of aES are partly known, the mechanism/s through which aES is regulated remains still elusive. In this review, we propose a model by which the small GTPase Rap1 could regulate aES assembly/remodelling. The characterization of key players in the dynamic of aES, such as Rap1, could open new possibility to develop prognostic, diagnostic, and therapeutic approaches for male patients under treatment for infertility as well as it could lead to the identification of new target for male contraception.

1. Introduction

Spermatogenesis is a very complex and regulated process during which the diploid spermatogonia divide and differentiate into haploid spermatozoa [14].

The correct development of fertile spermatozoa relies on the peculiar organization of the seminiferous epithelium. The germinal component (spermatogonia, primary and secondary spermatocytes, round spermatids, and elongating/elongated spermatids) is strictly interconnected with the somatic component, the Sertoli cells, which sustains spermatogenesis giving structural support and nourishment to germ cells [5, 6]. In mouse adult testis, the germ cells at different stages of differentiation display a unique pattern of association with Sertoli cells which can be classified into twelve stages (from I to XII) [1, 2, 4].

During these stages, spermatids undergo massive morphological modifications such as acquisition of cell polarity, condensation of chromatin, formation of the acrosome and tail, and production and elimination of the residual body. Meanwhile the differentiation process takes place, spermatids migrate across the entire length of the seminiferous epithelium until they reach the luminal edge where mature sperms are finally released [1, 7]. So, it is clear that germ cells have to remain anchored to Sertoli cells till the final steps in order to avoid a premature release as immature spermatids, with consequence on the male fertility potential (Figure 1).

The integrity of seminiferous epithelium and the functional cell interconnections are maintained through several junctional devices that take place between both the Sertoli-Sertoli cells and the Sertoli-germ cells. Besides junction types present also in other epithelia, like tight junctions [810] and gap junctions [11, 12], the seminiferous epithelium exhibits testis-unique anchoring junctions, as the ectoplasmic specialization (ES) [13, 14] and the desmosome-like junction [15].

The ES between the Sertoli cells is known as basal ES (bES). At the basal compartment (Figure 1), the bES coexists with other junctional structures like tight junction, gap junction, and desmosome-like junction; all together contribute to create the blood-testis barrier (BTB). The BTB physically divides the seminiferous epithelium in two compartments, that is, a basal compartment where spermatogonia and spermatocytes reside, and an adluminal compartment where spermatids differentiate to develop into spermatozoa (Figure 1) [16, 17]. The establishment of the BTB is fundamental for a successful spermatogenesis; its integrity has to be maintained throughout the entire spermatogenesis [18, 19]. The BTB provides in fact an immunological barrier to the developing male germ cells: it sequesters postmeiotic germ cells from the systemic circulation, thus preventing the production by the host of antibodies against spermatid-specific antigens whose expression is restricted to spermiogenesis only [20]. The BTB likely functions also as a gatekeeper, enabling only the passage through the seminiferous epithelium of selected substances/molecules of support to germ cells. The molecular components and the functions of the BTB have been extensively reviewed (for excellent reviews, see [12, 17, 21]); it will be no longer discussed here.

Conversely, this brief review will focus around the apical ES (aES), restricted to Sertoli-postmeiotic germ cells at the adluminal compartment. Differently from bES, aES does not coexist with other junctions: the aES is the only junctional device that sustains the association between Sertoli and elongating/elongated spermatids (from step 8 of differentiation) until the early phase of spermiation when it disassembles (Figure 1) [13, 2224].

2. Apical Ectoplasmic Specialization

The aES is the best known anchoring junction in the testis. Several studies have led to a significant improvement of our knowledge about its molecular architecture and molecular components. On the basis of such studies, the aES is emerged as a peculiar anchoring junction, being formed by structural components generally found in somatic adherens junctions (the cadherins/catenins and nectins/afadin complexes), tight junctions (such as Jam-C molecules), and focal contacts (the integrin/laminin complex) (Figure 2) [14, 25, 26]. This high heterogeneity is thought to permit that the aES accomplishes its multifunctional role in supporting spermiogenesis. In details, the aES makes the migration possible, concomitantly with the differentiation of spermatids across the seminiferous epithelium. Moreover, the aES could contribute to positioning elongating spermatids with their heads pointed towards the basal compartment. It is to notice that aES is first assembled within the seminiferous epithelium exactly when spermatids begin to loss their spherical shape to become a polarized cell (stage 8) [13, 14, 25]. Finally, the aES maintains spermatids attached to Sertoli cells until these are differentiated and ready to be released into the lumen [13, 23, 24]. It follows that aES has to undergo rapid cycles of assembly and disassembly, concomitantly with the progression of spermiogenesis, and that at spermiation the breakage at cell-cell contacts is definitive. The dynamic of these cycles must be finely and timely regulated. Despite many efforts, the mechanisms governing the aES remodeling during spermiogenesis/spermiation remain however obscure.

Herein, we address attention on how aES dynamic could be regulated. In particular, taking into consideration new experimental evidence provided independently from more laboratories, we discuss a model of regulation of aES assembly/disassembly upon the grounds of findings we obtained from an animal model that we generated appositely to inactivate the small Ras-like GTPase Rap1 [27]. Importantly, it is to underline that this model provides the first genetic link between Rap1 defects and male infertility; the Rap1[S17N] mutation is resulted to be instrumental in revealing the cAMP-Epac-Rap1/extracellular signal-regulated pathway that governs the spermatid-Sertoli cell adhesion at aES. Moreover, it is worth of mention to recall attention on the fact that the central cell of our model is the differentiating spermatid. So far, most works about the aES regulation highlight putative mechanisms operating in Sertoli cells, relegating spermatids to a role of merely spectators. However, the “actors” that are involved in cell-to-cell contact are at least two. Consequently, our model offers a new point of view to investigate about aES dynamic, that is, to consider that also germ cells could be actively engaged.

3. Rap1 Regulates aES Dynamic: Experimental Evidence

Rap1 is a member of the family of Ras-like small G proteins [28]; accordingly, it switches between an active conformation bound to the GTP and an inactive one bound to the GDP. The cycle between the two alternative states is coordinated by the guanine nucleotide exchange factors (GEFs), the activators that allow the binding with GTP, and the GTPase activating proteins (GAPs) that enhance the hydrolysis of bound GTP thus leading to Rap1 deactivation [29, 30]. Rap1 functions as a positional signal and organizer of cell architecture; it follows that this GTPase is placed upstream signalling pathways that regulate diverse cellular processes, including morphogenesis [31, 32], cell differentiation [33], cytoskeletal organization [34], cytokinesis [35], exocytosis/endocytosis [36], synaptic plasticity [37], and cell-cell adhesion [30]. As to this last aspect of Rap1 biology, a growing experimental evidence in the last years has allowed to highlight some aspects of Rap1 action in controlling integrin-based as well as cadherin-based junctional systems; in particular, Rap1 has been shown to regulate the levels of E-cadherin and VE-cadherin at the plasma membrane of epithelial and endothelial cells, respectively [3842].

Rap1 was first detected in the testis in 2000 [43]. It is expressed by germ cells throughout spermatogenesis; in spermatids, in particular, it was immunoprecipitated as a component of the signaling complex formed by the serine-threonine kinase B-Raf and the molecular adaptor 14-3-3 theta protein [43]. To verify the role of Rap1 in vivo in the process of sperm differentiation, we developed a mouse model. Transgenic mice that express a dominant negative mutant of Rap1 (iRap1) were generated; to have the expression of the mutant Rap1 variant selectively only in the postmeiotic germ cells, iRap1 was put under the control of the haploid-specific Protamine-1 promoter [27]. The phenotype of the mutant mice resulted in a derailment of spermiogenesis due to an anomalous release of immature round spermatids within the tubule lumen and low sperm counts. These findings addressed the research to point up towards the search of Rap1-regulated adhesion molecules leading to the discovery of VE-cadherin and of its epithelial cycle stage specific expression [27].

The high dynamicity of aES renders it as one of the most flexible cell-cell junctions in mammalian tissue, maybe comparable to the better characterized adherens junction at the endothelial cell barrier. This last is known to be a highly dynamic structure; it maintains the integrity of the endothelium, but it is also involved in the control of permeability, leukocyte diapedesis and, more generally speaking, vascular homeostasis [44]. The stabilization of endothelial adherens junctions relies on the stabilization at the plasma membrane of the vascular endothelial cadherin (VE-cadherin) through activation of Rap1 by the cAMP sensor/Rap1-GEF Epac [45]. It is, in fact, widely known that the increase of intracellular cAMP leads to the formation of circumferential actin bundles that support cadherins at adherens junctions; this occurs through Epac-activated Rap1. Similarly, Rap1 is involved in the formation of E-cadherin-based cell-cell adhesions in epithelial cells [38, 40, 41]. However, Aivatiadou et al. [27] found that not only VE-cadherin is expressed in the testis, but it localizes at aES level, exhibiting a pattern of expression that follows aES formation and function; it appears in the adluminal compartment when aESs are being formed and disappears at spermiation, with the exception of the intense staining of the soma of Sertoli cells at the basal compartment. Interestingly, in iRap mutant mice that express a dominant negative Rap1, VE-cadherin is more loosely linked to cytoskeleton and partially tyrosine phosphorylated, two conditions known to be related to impairment of cell-cell adhesions [27, 41]. At last, we remember that the adaptor proteins -, -catenins, essential for linking the cadherins to actin filaments [46], and p120-catenin are expressed in both Sertoli and germ cells [47]. Similarly, germ cells express also afadin [48]. Afadin is a scaffold protein containing an actin-binding and Ras/Rap-binding domain [29, 49] that has been reported to regulate the cyclical activation/inactivation of Rap1 and RhoA [50]. In endothelial and epithelial cells, afadin is involved in Rap1-dependent assembly of cadherins-based adherens junctions (AJs) [40, 51, 52]. The Rap1-GTP/afadin complex mediates the recruitment of p120-catenin to the plasma membrane [51, 52], thus stabilizing VE-cadherin and protecting it from endocytosis [53].

The iRap1 animal model by Aivatiadou et al. [27] clearly demonstrated that Rap1 is involved in the regulation of aES junction dynamic. However, how Rap1 exerts its control and what are its protein targets have not been yet dissected. Very likely, testis VE-cadherin is the adhesion receptor target of Rap1 in spermatids; this, however, does not mean that VE-cadherin is the only one. Here, we suggest, in addition to VE-cadherin, another putative candidate, nectin. Nectins belong to the superfamily of Ca2+-independent immunoglobulins that comprises at least four members [49]; they are able to form both homophilic and heterophilic trans-interactions [49]. Nectins have been shown at aESs; more specifically, nectin-2 on the Sertoli membrane trans-interacts with nectin-3 on the spermatid membrane [54]. It has been reported that the nectin-2/nectin-3 complex at aES is stabilized by afadin that connects the complex to the actin filaments [54]. Since afadin could connect with both nectin-based and cadherin-based junctional systems, Rap1 could represent the central regulatory link between these two aES structural complexes. Considering that aESs undergo cycles of assembly and disassembly and that Rap1 mediates both the de novo formation and the reestablishment of adherens junctions [38, 55], the following hypothesis for future experimental research could be suggested. That is, at the assembling, nectins are first engaged at aES recruiting afadin that, on its own, is involved in activation of Rap1; this leads to an accumulation of VE-cadherin at the plasma membrane with the result of strengthening of the junction. The nectins/VE-cadherin adhesion molecules have already been described to be able to connect physically and functionally in endothelial cell systems [56, 57].

4. Rap1 as an Organizer of Spermatid Polarization at aES

Rap1 action is accomplished through the cooperation of different signaling pathways involving several effectors [32, 58, 59]. In its central role of positional signal and organizer of cell architecture, Rap1 governs the reorganization of actin cytoskeleton [34, 60, 61]. The great plasticity of the aES requires a rapid rearrangement of actin cytoskeleton; it is not surprising if Rap1 could emerge as a regulator of this cytoskeleton at aES. CDC42 is another Ras-like small GTPase that belongs to the family of Rho-GTPases [6264], and CDC42 is a Rap1 effector. In epithelial as well as endothelial cells, Epac-activated Rap1 induces CDC42 activation; this leads to a reorganization of actin cytoskeleton resulting in the formation of circumferential actin bundles and consequently in the stability of E-cadherin/VE-cadherin-based cell-cell adhesions [34, 45, 60]. Since CDC42 is expressed in differentiating spermatids [65, 66], it is possible to speculate that Rap1-mediated CDC42 activation is one of the mechanisms that stabilize cell-cell contacts at aESs. Not only, but CDC42 may be the Rap1 effector through which Rap1, acting as a critical regulator of cell polarization [67], controls spermatid polarization, an event for which the contribution of aES has been evoked more times, in the seminiferous epithelium. Cdc42, in fact, is known to work in epithelial cells in concert with the Par-based polarity protein complex to establish the apicobasal cell polarity [68, 69]. Interestingly, in spermatids the Par6/Cdc42/aPKC complex has been shown to be recruited to the plasma membrane by Jam-C, a junctional adhesion molecule found at aES level [65, 70]; Gliki et al. [65] attributed to Jam-C the role of assembling the cell polarity complex and, consequently, of promoting spermatid polarization. However, the pathway though which Jam-C could fulfill such functions was not dissected. In endothelial cells, Jam-C regulates vascular endothelial permeability by modulating VE-cadherin-mediated cell-cell contacts [71]. In particular, the loss of Jam-C expression by Jam-C knockdown results in stabilization of VE-cadherin-mediated adhesion in a Rap1-dependent manner. It follows that the question of Jam-C and Par6/Cdc42/aPKC complex in differentiating spermatids deserves a deeper investigation. Again, Rap1 may emerge as the central regulator that supervises both aES assembly/disassembly and spermatid polarity in a new system, the spermatid-Sertoli cell adhesion system. In this regard, it is to notice that just the VE-cadherin-based AJs have been reported to be essential to establish cell polarity; here, it is in fact recruited the Rap1-activated cell polarity complex in endothelia [72]. So, it is not to exclude that Rap1 could mediate spermatid polarization not only through the Jam-C/polarity complex but also through the VE-cadherin/polarity complex system.

5. Rap1, Rho, and the Disassembling of aES

Rho belongs, as CDC42 and Rac1, to the family of Rho GTPases and is involved in the dynamics of F-actin structures thus influencing cell shape and assembly of AJs. The best characterized Rho isoforms are RhoA, RhoB, and RhoC [73, 74]; RhoA and RhoB have been reported in testis and spermatozoa [66]. While CDC42 and Rac1 are known to act as Rap1 effectors, that is, downstream the activated Rap1, Rho GTPase is best known to counteract the action of Rap1 [27, 34, 75]. For example, activation of the Rho GTPase pathway determines endothelial cell hyperpermeability and could lead to endothelial barrier disruption [75, 76]. Accordingly, overexpression of a constitutively active mutant of Rap1 results in activation of Rac1 and, intriguingly, inactivation of RhoA [50]. As already noticed, with few exceptions, members of the Ras-like GTPase super-family cycle, between an active and inactive state. The continuous cycling between the two states is tightly controlled by a number of regulatory proteins that specify where and for how long the signal is “on” and which cellular function is modulated. In the case of cell adhesion, for each cellular process triggered by cell-cell contacts, multiple GTPases must be dynamically turned “on” or “off” [34, 50, 75]. This “turning” is facilitated by GEFs and GAPs, respectively. So far, several GEFs and GAPs have been described to modulate the organization, molecular composition, and function of adhesive complexes in different cell types. In endothelial cells, the second messenger cAMP induces barrier protective responses against thrombin or inflammatory mediators [77, 78]. The Rap1 GEFs that are sensors of cAMP elevation are known as Epac (exchange protein activated by cAMP), of which there are two variants, Epac1 and Epac2 [7981]. Rap1, however, could be activated also by other GEFs, such as C3G, PDZ-GEFs, and CalDAGs [59, 82, 83]. Still referring to the endothelial cell AJs, Birukova et al. [75] have shown that the Rap1 PDZ-GEF cooperates with Epac to maintain junction integrity; more specifically, the PDZ-GEF is involved mainly in Rap1 activation under resting conditions [45], while Epac/s is/are necessary to further tighten cell-cell contacts [45]. Both Epac1 and Epac2 have been found in mouse testis and male germ cells [80, 84]. Similar to the GEFs, there are various Rap-GAPs that are specifically targeted to different molecular complexes at various cellular locations [59]. These GAPs may reverse the dynamic processes controlled by activated Rap1. Among the best characterized Rap1-GAPs, there are Rap1-GAP1,2, Spa-1, and SPAR1,2,3 [59]; so far, however, experimental evidence for Rap1-GAPs in the testis is still lacking.

This is not the case, on the contrary, for RhoGAPs. Aivatiadou et al. [80] reported that male germ cells express RA-RhoGAP. RA-RhoGAP is a RhoGAP that possesses a RA domain through which it binds to Epac-activated Rap1 thus acting as a Rap1 effector and transductor of signal from Rap1 to Rho [85]. Indeed, Aivatiadou et al. [80] showed also that in isolated spermatogenic cells, after stimulation with 8-CPT (a membrane-permeable analogue of cAMP), Epac-activated Rap1 colocalizes with RA-RhoGAP. It follows that male germ cells have the equipment of signaling molecules, including activators and effectors, through which activated Rap1 could suppress the action of Rho.

At this point, if Rap1 and Rho are likely the Ras-like GTPases that govern aES dynamic, a key question concerns the signal/s responsible for the Rap1/Rho activation/deactivation at aES. Before the conclusion of this brief review, we want to provide a further suggestion towards the direction for this putative signaling molecule. Transforming growth factor-1 (TGF-1) functions in diverse cellular processes, such as tissue differentiation and cell migration. Recent experimental evidence on signaling that governs monocyte adhesion and chemotaxis [86] has revealed that TGF-1 triggers cAMP elevation leading to Rap1 activation via Epac; not only this, but also this Rap1 activation, on its own, results in Rho inactivation through the Rap1-dependent RhoGAP. In other words, prolonged TGF-1-treated cells produce cAMP, which activates sequentially Epac, Rap1, and Rap1-dependent RhoGAP, resulting in suppression of Rho and macrophage migration. TGF-1 is produced in the testis by both germ cells and Sertoli cells [87]. TGF-1 may thus result to be crucial for the restructuring of aES.

Conclusively, here we provide a model (Figure 3) for the mechanism/s by which Rap1 could potentially regulate aES dynamic. It is to remark that here we took into consideration deliberately only spermatids, that is, the cells so far rather neglected under this context. Restructuring of aES junction is thought to be dependent on a highly coordinated network of mechanisms of activation/deactivation of Ras-like GTPases that has Rap1 as its central hub and the Rho-family GTPases as downstream Rap1-effectors/antagonists. Rap1 action results in the assembly/stabilization/reestablishment of aES junctions whereas Rho drives their disassembling; interestingly, the functional Rap1/Rho interaction is proposed to be vice-versa reciprocal. Obviously, some aspects of the suggested regulation of aES dynamic need to be verified and are waiting for the experimental validation.

6. Conclusion

Male infertility is one of the health problems worldwide. Several defects responsible for male infertility are still unclear, mostly due to our poor understanding of molecular mechanisms that regulate sperm production and release from the testis, the maturation and transit of the sperm through the male and female tracts, and the events essential for fertilization. Because we do not understand entirely these molecular processes, we cannot diagnose correctly the causes of male infertility. Upon the grounds of the phenotype that characterizes the unique animal model developed so far to investigate in vivo about the nature of the signaling involved in the regulation of spermatid-Sertoli cell junctions [27], here we propose two interconnected mechanisms that could unravel the question of the regulation of a highly dynamic and essential junction like the aES is.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


  1. C. P. Leblond and Y. Clermont, “Definition of the stages of the cycle of the seminiferous epithelium in the rat,” Annals of the New York Academy of Sciences, vol. 55, no. 4, pp. 548–573, 1952. View at: Google Scholar
  2. E. F. Oakberg, “A description of spermiogenesis in the mouse and its use in analysis of the cycle of the seminiferous epithelium and germ cell renewal,” The American Journal of Anatomy, vol. 99, no. 3, pp. 391–413, 1956. View at: Google Scholar
  3. L. Russell, R. A. Ettlin, A. P. Sinha Hikim, and E. J. Clegg, Histological and Histopathological Evaluation of the Testis, Cache River Press, Clearwater, Fla, USA, 1990.
  4. R. A. Hess and L. R. de Franca, “Spermatogenesis and cycle of the seminiferous epithelium,” Advances in Experimental Medicine and Biology, vol. 636, pp. 1–15, 2008. View at: Publisher Site | Google Scholar
  5. M. D. Griswold, “The central role of Sertoli cells in spermatogenesis,” Seminars in Cell and Developmental Biology, vol. 9, no. 4, pp. 411–416, 1998. View at: Google Scholar
  6. M. Godet, O. Sabido, J. Gilleron, and P. Durand, “Meiotic progression of rat spermatocytes requires mitogen-activated protein kinases of Sertoli cells and close contacts between the germ cells and the Sertoli cells,” Developmental Biology, vol. 315, no. 1, pp. 173–188, 2008. View at: Publisher Site | Google Scholar
  7. L. Russell, “Movement of spermatocytes from the basal to the adluminal compartment of the rat testis,” The American Journal of Anatomy, vol. 148, no. 3, pp. 313–328, 1977. View at: Google Scholar
  8. L. D. Russell and R. N. Peterson, “Sertoli cell junctions: morphological and functional correlates,” International Review of Cytology, vol. 94, pp. 177–211, 1985. View at: Google Scholar
  9. D. D. Mruk and C. Y. Cheng, “Sertoli-Sertoli and Sertoli-germ cell interactions and their significance in germ cell movement in the seminiferous epithelium during spermatogenesis,” Endocrine Reviews, vol. 25, no. 5, pp. 747–806, 2004. View at: Publisher Site | Google Scholar
  10. L. Su, D. D. Mruk, and C. Y. Cheng, “Regulation of the blood-testis barrier by coxsackievirus and adenovirus receptor,” The American Journal of Physiology, vol. 303, no. 8, pp. C843–C853, 2012. View at: Google Scholar
  11. K. Weider, M. Bergmann, and R. Brehm, “Connexin 43: its regulatory role in testicular junction dynamics and spermatogenesis,” Histology and Histopathology, vol. 26, no. 10, pp. 1343–1352, 2011. View at: Google Scholar
  12. C. Y. Cheng, E. W. Wong, P. P. Lie, M. W. Li, D. D. Mruk, H. H. Yan et al., “Regulation of blood-testis barrier dynamics by desmosome, gap junction, hemidesmosome and polarity proteins: an unexpected turn of events,” Spermatogenesis, vol. 1, no. 2, pp. 105–115, 2011. View at: Google Scholar
  13. A. W. Vogl, D. C. Pfeiffer, D. Mulholland, G. Klmel, and J. Guttman, “Unique and multifunctional adhesion junctions in the testis: ectoplasmic specializations,” Archives of Histology and Cytology, vol. 63, no. 1, pp. 1–15, 2000. View at: Google Scholar
  14. E. W. P. Wong, D. D. Mruk, and C. Y. Cheng, “Biology and regulation of ectoplasmic specialization, an atypical adherens junction type, in the testis,” Biochimica et Biophysica Acta, vol. 1778, no. 3, pp. 692–708, 2008. View at: Publisher Site | Google Scholar
  15. D. D. Mruk and C. Y. Cheng, “Desmosomes in the testis: moving into an unchartered territory,” Spermatogenesis, vol. 1, no. 1, pp. 47–51, 2011. View at: Google Scholar
  16. M. Dym and D. W. Fawcett, “The blood-testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium,” Biology of Reproduction, vol. 3, no. 3, pp. 308–326, 1970. View at: Google Scholar
  17. C.-H. Wong and C. Y. Cheng, “The blood-testis barrier: its biology, regulation, and physiological role in spermatogenesis,” Current Topics in Developmental Biology, vol. 71, pp. 263–296, 2005. View at: Publisher Site | Google Scholar
  18. J. Grima and C. Y. Cheng, “Testin induction: the role of cyclic 3′,5′-adenosine monophosphate/protein kinase A signaling in the regulation of basal and lonidamine-induced testin expression by rat Sertoli cells,” Biology of Reproduction, vol. 63, no. 6, pp. 1648–1660, 2000. View at: Google Scholar
  19. N. P. Y. Chung and C. Y. Cheng, “Is cadmium chloride-induced inter-Sertoli tight junction permeability barrier disruption a suitable in vitro model to study the events of junction disassembly during spermatogenesis in the rat testis?” Endocrinology, vol. 142, no. 5, pp. 1878–1888, 2001. View at: Publisher Site | Google Scholar
  20. K. W. Beagley, Z. L. Wu, M. Pomering, and R. C. Jones, “Immune responses in the epididymis: implications for immunocontraception,” Journal of Reproduction and Fertility, vol. 53, pp. 235–245, 1998. View at: Google Scholar
  21. C. Yan Cheng and D. D. Mruk, “The blood-testis barrier and its implications for male contraception,” Pharmacological Reviews, vol. 64, no. 1, pp. 16–64, 2012. View at: Publisher Site | Google Scholar
  22. L. Russell, “Observations on rat Sertoli ectoplasmic (junctional) specializations in their association with germ cells of the rat testis,” Tissue and Cell, vol. 9, no. 3, pp. 475–498, 1977. View at: Google Scholar
  23. L. D. Russell, “Morphological and functional evidence for Sertoli-germ cell relationships,” in The Sertoli Cell, L. D. Russell and M. D. Griswold, Eds., pp. 365–390, Cache River Press, Clearwater, Fla, USA, 1993. View at: Google Scholar
  24. J. S. Young, J. A. Guttman, K. S. Vaid, and A. Wayne Vogl, “Tubulobulbar complexes are intercellular podosome-like structures that internalize intact intercellular junctions during epithelial remodeling events in the rat testis,” Biology of Reproduction, vol. 80, no. 1, pp. 162–174, 2009. View at: Publisher Site | Google Scholar
  25. D. D. Mruk, B. Silvestrini, and C. Y. Cheng, “Anchoring junctions as drug targets: role in contraceptive development,” Pharmacological Reviews, vol. 60, no. 2, pp. 146–180, 2008. View at: Publisher Site | Google Scholar
  26. I. A. Ropera, B. Bilinska, C. Y. Cheng, and D. D. Mruk, “Sertoli-germ cell junctions in the testis: a review of recent data,” Philosophical Transactions of the Royal Society B, vol. 365, no. 1546, pp. 1593–1605, 2010. View at: Publisher Site | Google Scholar
  27. E. Aivatiadou, E. Mattei, M. Ceriani, L. Tilia, and G. Berruti, “Impaired fertility and spermiogenetic disorders with loss of cell adhesion in male mice expressing an interfering Rap1 mutant,” Molecular Biology of the Cell, vol. 18, no. 4, pp. 1530–1542, 2007. View at: Publisher Site | Google Scholar
  28. H. Kitayama, Y. Sugimoto, T. Matsuzaki, Y. Ikawa, and M. Noda, “A ras-related gene with transformation suppressor activity,” Cell, vol. 56, no. 1, pp. 77–84, 1989. View at: Google Scholar
  29. M. R. H. Kooistra, N. Dubé, and J. L. Bos, “Rap1: a key regulator in cell-cell junction formation,” Journal of Cell Science, vol. 120, no. 1, pp. 17–22, 2007. View at: Publisher Site | Google Scholar
  30. W.-J. Pannekoek, M. R. H. Kooistra, F. J. T. Zwartkruis, and J. L. Bos, “Cell-cell junction formation: the role of Rap1 and Rap1 guanine nucleotide exchange factors,” Biochimica et Biophysica Acta, vol. 1788, no. 4, pp. 790–796, 2009. View at: Publisher Site | Google Scholar
  31. M. Ji and O. M. Andrisani, “High-level activation of cyclic AMP signaling attenuates bone morphogenetic protein 2-induced sympathoadrenal lineage development and promotes melanogenesis in neural crest cultures,” Molecular and Cellular Biology, vol. 25, no. 12, pp. 5134–5145, 2005. View at: Publisher Site | Google Scholar
  32. M. Chrzanowska-Wodnicka, “Distinct functions for Rap1 signaling in vascular morphogenesis and dysfunction,” Experimental Cell Research, vol. 319, no. 15, pp. 2350–2359, 2013. View at: Google Scholar
  33. L. Li, J. S. Kim, and V. A. Boussiotis, “Rap1A regulates generation of T regulatory cells via LFA-1-dependent and LFA-1-independent mechanisms,” Cellular Immunology, vol. 266, no. 1, pp. 7–13, 2010. View at: Publisher Site | Google Scholar
  34. K. Ando, S. Fukuhara, T. Moriya, Y. Obara, N. Nakahata, and N. Mochizuki, “Rap1 potentiates endothelial cell junctions by spatially controlling myosin II activity and actin organization,” Journal of Cell Biology, vol. 202, no. 6, pp. 901–916, 2013. View at: Google Scholar
  35. V. T. Dao, A. G. Dupuy, O. Gavet, E. Caron, and J. de Gunzburg, “Dynamic changes in Rap1 activity are required for cell retraction and spreading during mitosis,” Journal of Cell Science, vol. 122, no. 16, pp. 2996–3004, 2009. View at: Publisher Site | Google Scholar
  36. K. W. van Hooren, E. L. van Agtmaal, M. Fernandez-Borja, J. A. van Mourik, J. Voorberg, and R. Bierings, “The Epac-Rap1 signaling pathway controls cAMP-mediated exocytosis of Weibel-Palade bodies in endothelial cells,” Journal of Biological Chemistry, vol. 287, no. 29, pp. 24713–24720, 2012. View at: Google Scholar
  37. R. L. Stornetta and J. J. Zhu, “Ras and Rap signaling in synaptic plasticity and mental disorders,” Neuroscientist, vol. 17, no. 1, pp. 54–78, 2011. View at: Publisher Site | Google Scholar
  38. C. Hogan, N. Serpente, P. Cogram et al., “Rap1 regulates the formation of E-cadherin-based cell-cell contacts,” Molecular and Cellular Biology, vol. 24, no. 15, pp. 6690–6700, 2004. View at: Publisher Site | Google Scholar
  39. T. Hoshino, T. Sakisaka, T. Baba, T. Yamada, T. Kimura, and Y. Takai, “Regulation of E-cadherin endocytosis by nectin through afadin, Rap1, and p120ctn,” Journal of Biological Chemistry, vol. 280, no. 25, pp. 24095–24103, 2005. View at: Publisher Site | Google Scholar
  40. T. Sato, N. Fujita, A. Yamada et al., “Regulation of the assembly and adhesion activity of E-cadherin by nectin and afadin for the formation of adherens junctions in Madin-Darby canine kidney cells,” Journal of Biological Chemistry, vol. 281, no. 8, pp. 5288–5299, 2006. View at: Publisher Site | Google Scholar
  41. S. Fukuhara, A. Sakurai, A. Yamagishi, K. Sako, and N. Moehizuki, “Vascular endothelial cadherin-mediated cell-cell adhesion regulated by a small GTPase, Rap1,” Journal of Biochemistry and Molecular Biology, vol. 39, no. 2, pp. 132–139, 2006. View at: Google Scholar
  42. K. Noda, J. Zhang, S. Fukuhara, S. Kunimoto, M. Yoshimura, and N. Mochizuki, “Vascular endothelial-cadherin stabilizes at cell-cell junctions by anchoring to circumferential actin bundles through α- and β-catenins in cyclic AMP-Epac-Rap1 signal-activated endothelial cells,” Molecular Biology of the Cell, vol. 21, no. 4, pp. 584–596, 2010. View at: Publisher Site | Google Scholar
  43. G. Berruti, “A novel Rap1/B-Raf/14-3-3 θ protein complex is formed in vivo during the morphogenetic differentiation of postmeiotic male germ cells,” Experimental Cell Research, vol. 257, no. 1, pp. 172–179, 2000. View at: Publisher Site | Google Scholar
  44. E. Dejana, “Endothelial cell-cell junctions: happy together,” Nature Reviews Molecular Cell Biology, vol. 5, no. 4, pp. 261–270, 2004. View at: Publisher Site | Google Scholar
  45. W.-J. Pannekoek, J. J. G. van Dijk, O. Y. A. Chan et al., “Epac1 and PDZ-GEF cooperate in Rap1 mediated endothelial junction control,” Cellular Signalling, vol. 23, no. 12, pp. 2056–2064, 2011. View at: Publisher Site | Google Scholar
  46. C. Kintner, “Regulation of embryonic cell adhesion by the cadherin cytoplasmic domain,” Cell, vol. 69, no. 2, pp. 225–236, 1992. View at: Publisher Site | Google Scholar
  47. N. P. Y. Lee, D. Mruk, W. M. Lee, and C. Y. Cheng, “Is the cadherin/catenin complex a functional unit of cell-cell actin-based adherens junctions in the rat testis?” Biology of Reproduction, vol. 68, no. 2, pp. 489–508, 2003. View at: Publisher Site | Google Scholar
  48. H. H. N. Yan, D. D. Mruk, W. M. Lee, and C. Y. Cheng, “Ectoplasmic specialization: a friend or a foe of spermatogenesis?” BioEssays, vol. 29, no. 1, pp. 36–48, 2007. View at: Publisher Site | Google Scholar
  49. Y. Takai, W. Ikeda, H. Ogita, and Y. Rikitake, “The immunoglobulin-like cell adhesion molecule nectin and its associated protein afadin,” Annual Review of Cell and Developmental Biology, vol. 24, pp. 309–342, 2008. View at: Publisher Site | Google Scholar
  50. M. Miyata, Y. Rikitake, M. Takahashi et al., “Regulation by afadin of cyclical activation and inactivation of Rap1, Rac1, and RhoA small G proteins at leading edges of moving NIH3T3 cells,” Journal of Biological Chemistry, vol. 284, no. 36, pp. 24595–24609, 2009. View at: Publisher Site | Google Scholar
  51. H. Tawa, Y. Rikitake, M. Takahashi et al., “Role of afadin in vascular endothelial growth factor-and sphingosine 1-phosphate-induced angiogenesis,” Circulation Research, vol. 106, no. 11, pp. 1731–1742, 2010. View at: Publisher Site | Google Scholar
  52. A. A. Birukova, P. Fu, T. Wu et al., “Afadin controls p120-catenin-ZO-1 interactions leading to endothelial barrier enhancement by oxidized phospholipids,” Journal of Cellular Physiology, vol. 227, no. 5, pp. 1883–1890, 2012. View at: Publisher Site | Google Scholar
  53. K. Xiao, J. Garner, K. M. Buckley et al., “p120-catenin regulates clathrin-dependent endocytosis of VE-cadherin,” Molecular Biology of the Cell, vol. 16, no. 11, pp. 5141–5151, 2005. View at: Publisher Site | Google Scholar
  54. K. Ozaki-Kuroda, H. Nakanishi, H. Ohta et al., “Nectin couples cell-cell adhesion and the actin scaffold at heterotypic testicular junctions,” Current Biology, vol. 12, no. 13, pp. 1145–1150, 2002. View at: Publisher Site | Google Scholar
  55. E. S. Wittchen, R. A. Worthylake, P. Kelly, P. J. Casey, L. A. Quilliam, and K. Burridge, “Rap1 GTPase inhibits leukocyte transmigration by promoting endothelial barrier function,” Journal of Biological Chemistry, vol. 280, no. 12, pp. 11675–11682, 2005. View at: Publisher Site | Google Scholar
  56. T. Fukuyama, H. Ogita, T. Kawakatsu et al., “Involvement of the c-Src-Crk-C3G-Rap1 signaling in the nectin-induced activation of Cdc42 and formation of adherens junctions,” Journal of Biological Chemistry, vol. 280, no. 1, pp. 815–825, 2005. View at: Publisher Site | Google Scholar
  57. D. Herr, H. M. Fraser, R. Konrad, I. Holzheu, R. Kreienberg, and C. Wulff, “Human chorionic gonadotropin controls luteal vascular permeability via vascular endothelial growth factor by down-regulation of a cascade of adhesion proteins,” Fertility and Sterility, vol. 99, no. 6, pp. 1749–1758, 2013. View at: Google Scholar
  58. S. Citi, D. Spadaro, Y. Schneider, J. Stutz, and P. Pulimeno, “Regulation of small GTPases at epithelial cell-cell junctions,” Molecular Membrane Biology, vol. 28, no. 7-8, pp. 427–444, 2011. View at: Publisher Site | Google Scholar
  59. M. Gloerich and J. L. Bos, “Regulating Rap small G-proteins in time and space,” Trends in Cell Biology, vol. 21, no. 10, pp. 615–623, 2011. View at: Publisher Site | Google Scholar
  60. M. R. H. Kooistra, M. Corada, E. Dejana, and J. L. Bos, “Epac1 regulates integrity of endothelial cell junctions through VE-cadherin,” FEBS Letters, vol. 579, no. 22, pp. 4966–4972, 2005. View at: Publisher Site | Google Scholar
  61. A. A. Birukova, D. Burdette, N. Moldobaeva, J. Xing, P. Fu, and K. G. Birukov, “Rac GTPase is a hub for protein kinase A and Epac signaling in endothelial barrier protection by cAMP,” Microvascular Research, vol. 79, no. 2, pp. 128–138, 2010. View at: Publisher Site | Google Scholar
  62. A. B. Jaffe and A. Hall, “Rho GTPases: biochemistry and biology,” Annual Review of Cell and Developmental Biology, vol. 21, pp. 247–269, 2005. View at: Publisher Site | Google Scholar
  63. K. Wennerberg and C. J. Der, “Rho-family GTPases: it's not only Rac and Rho (and I like it),” Journal of Cell Science, vol. 117, no. 8, pp. 1301–1312, 2004. View at: Publisher Site | Google Scholar
  64. J. Melendez, M. Grogg, and Y. Zheng, “Signaling role of Cdc42 in regulating mammalian physiology,” Journal of Biological Chemistry, vol. 286, no. 4, pp. 2375–2381, 2011. View at: Publisher Site | Google Scholar
  65. G. Gliki, K. Ebnet, M. Aurrand-Lions, B. A. Imhof, and R. H. Adams, “Spermatid differentiation requires the assembly of a cell polarity complex downstream of junctional adhesion molecule-C,” Nature, vol. 431, no. 7006, pp. 320–324, 2004. View at: Publisher Site | Google Scholar
  66. R. Baltiérrez-Hoyos, A. L. Roa-Espitia, and E. O. Hernández-González, “The association between CDC42 and caveolin-1 is involved in the regulation of capacitation and acrosome reaction of guinea pig and mouse sperm,” Reproduction, vol. 144, no. 1, pp. 123–134, 2012. View at: Google Scholar
  67. Y. Jossin, “Polarization of migrating cortical neurons by Rap1 and N-cadherin, revisiting the model for the reelin signaling pathway,” Small GTPases, vol. 2, no. 6, pp. 322–328, 2011. View at: Google Scholar
  68. D. P. Welchman, L. D. Mathies, and J. Ahringer, “Similar requirements for CDC-42 and the PAR-3/PAR-6/PKC-3 complex in diverse cell types,” Developmental Biology, vol. 305, no. 1, pp. 347–357, 2007. View at: Publisher Site | Google Scholar
  69. K. P. Harris and U. Tepass, “Cdc42 and vesicle trafficking in polarized cells,” Traffic, vol. 11, no. 10, pp. 1272–1279, 2010. View at: Publisher Site | Google Scholar
  70. E. W. P. Wong, D. D. Mruk, W. M. Lee, and C. Y. Cheng, “Par3/Par6 polarity complex coordinates apical ectoplasmic specialization and blood-testis barrier restructuring during spermatogenesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 28, pp. 9657–9662, 2008. View at: Publisher Site | Google Scholar
  71. V. V. Orlova, M. Economopoulou, F. Lupu, S. Santoso, and T. Chavakis, “Junctional adhesion molecule-C regulates vascular endothelial permeability by modulating VE-cadherin-mediated cell-cell contacts,” Journal of Experimental Medicine, vol. 203, no. 12, pp. 2703–2714, 2006. View at: Publisher Site | Google Scholar
  72. M. G. Lampugnani, F. Orsenigo, N. Rudini et al., “CCM1 regulates vascular-lumen organization by inducing endothelial polarity,” Journal of Cell Science, vol. 123, no. 7, pp. 1073–1080, 2010. View at: Publisher Site | Google Scholar
  73. C. L. Hutchinson, P. N. Lowe, S. H. McLaughlin, H. R. Mott, and D. Owen, “Differential binding of RhoA, RhoB, and RhoC to protein kinase C-related kinase (PRK) isoforms PRK1, PRK2, and PRK3: PRKs have the highest affinity for RhoB,” Biochemistry, vol. 52, no. 45, pp. 7999–8011, 2013. View at: Google Scholar
  74. A. J. Ridley, “RhoA, RhoB and RhoC have different roles in cancer cell migration,” Journal of Microscopy, vol. 251, no. 3, pp. 242–249, 2013. View at: Google Scholar
  75. A. A. Birukova, X. Tian, Y. Tian, K. Higginbotham, and K. G. Birukova, “Rap-afadin axis in control of Rho signaling and endothelial barrier recovery,” Molecular Biology of the Cell, vol. 24, no. 17, pp. 2678–2688, 2013. View at: Google Scholar
  76. X. Cullere, S. K. Shaw, L. Andersson, J. Hirahashi, F. W. Luscinskas, and T. N. Mayadas, “Regulation of vascular endothelial barrier function by Epac, a cAMP-activated exchange factor for Rap GTPase,” Blood, vol. 105, no. 5, pp. 1950–1955, 2005. View at: Publisher Site | Google Scholar
  77. A. A. Birukova, T. Zagranichnaya, P. Fu et al., “Prostaglandins PGE2 and PGI2 promote endothelial barrier enhancement via PKA- and Epac1/Rap1-dependent Rac activation,” Experimental Cell Research, vol. 313, no. 11, pp. 2504–2520, 2007. View at: Publisher Site | Google Scholar
  78. N. V. Bogatcheva, J. G. Garcia, and A. D. Verin, “Molecular mechanisms of thrombin-induced endothelial cell permeability,” Biochemistry, vol. 67, no. 1, pp. 75–84, 2002. View at: Google Scholar
  79. H. Kawasaki, G. M. Springett, N. Mochizuki et al., “A family of cAMP-binding proteins that directly activate Rap1,” Science, vol. 282, no. 5397, pp. 2275–2279, 1998. View at: Publisher Site | Google Scholar
  80. E. Aivatiadou, M. Ripolone, F. Brunetti, and G. Berruti, “cAMP-Epac2-mediated activation of Rap1 in developing male germ cells: RA-RhoGAP as a possible direct down-stream effector,” Molecular Reproduction and Development, vol. 76, no. 4, pp. 407–416, 2009. View at: Publisher Site | Google Scholar
  81. M. Gloerich and J. L. Bos, “Epac: defining a new mechanism for cAMP action,” Annual Review of Pharmacology and Toxicology, vol. 50, pp. 355–375, 2010. View at: Publisher Site | Google Scholar
  82. S. Asuri, J. Yan, N. C. Paranavitana, and L. A. Quilliam, “E-cadherin dis-engagement activates the Rap1 GTPase,” Journal of Cellular Biochemistry, vol. 105, no. 4, pp. 1027–1037, 2008. View at: Publisher Site | Google Scholar
  83. G. F. Guidetti, D. Manganaro, A. Consonni, I. Canobbio, C. Balduini, and M. Torti, “Phosphorylation of the guanine-nucleotide-exchange factor CalDAG-GEFI by protein kinase A regulates Ca(2+)-dependent activation of platelet Rap1b GTPase,” Biochemical Journal, vol. 453, no. 1, pp. 115–123, 2013. View at: Google Scholar
  84. G. Berruti, “cAMP activates Rap1 in differentiating mouse male germ cells: a new signaling pathway mediated by the cAMP-activated exchange factor Epac?” Cellular and Molecular Biology, vol. 49, no. 3, pp. 381–388, 2003. View at: Google Scholar
  85. T. Yamada, T. Sakisaka, S. Hisata, T. Baba, and Y. Takai, “RA-RhoGAP, Rap-activated Rho GTPase-activating protein implicated in neurite outgrowth through Rho,” Journal of Biological Chemistry, vol. 280, no. 38, pp. 33026–33034, 2005. View at: Publisher Site | Google Scholar
  86. M. Y. Moon, H. J. Kim, J. G. Kim, J. Y. Lee, J. Kim et al., “Small GTPase Rap1 regulates cell migration through regulation of small GTPase RhoA activity in response to transforming growth factor-β1,” Journal of Cellular Physiology, vol. 228, no. 11, pp. 2119–2126, 2013. View at: Google Scholar
  87. K. J. Teerds and J. H. Dorrington, “Localization of transforming growth factor β1 and β2 during testicular development in the rat,” Biology of Reproduction, vol. 48, no. 1, pp. 40–45, 1993. View at: Publisher Site | Google Scholar

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