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The apical plasma membrane of Drosophila embryonic epithelia
ARTICLE IN PRESS European Journal of Cell Biology 89 (2010) 208–211
Contents lists available at ScienceDirect
European Journal of Cell Biology journal homepage: www.elsevier.de/ejcb
The apical plasma membrane of Drosophila embryonic epithelia Anne Uv a, Bernard Moussian b,n a b
Institute of Biomedicine, Department of Medicinal Genetics, Gothenburg University, Medicinaregatan 9A, 40530 Gothenburg, Sweden Max-Planck Institute for Developmental Biology, Spemannstr. 35, 72076 T¨ ubingen, Germany
a r t i c l e in f o
Keywords: Epithelia Epidermis Trachea Midgut Apical plasma membrane Microvilli Cuticle Taenidia Apical undulae
a b s t r a c t The apical plasma membrane of epithelia presents the interface between organs and the external environment. It has biochemical activities distinct from those of the basal and lateral plasma membranes, as it accommodates the production and assembly of ordered apical matrices involved in organ protection and physiology and determines the microenvironment in the apical extracellular milieu. Here, we emphasise the importance of the apical plasma membrane in tissue differentiation, by mainly focussing on the embryo of the fruit fly Drosophila melanogaster, and discuss the principal organisation of the apical plasma membrane into repetitive subdomains of specific topologies and activities essential for epithelial function. & 2009 Elsevier GmbH. All rights reserved.
Introduction The plasma membrane of epithelial cells is subdivided into three functional areas, the apical, the lateral and the basal plasma membrane (Fig. 1) (Tepass and Hartenstein, 1994), that are defined by distinct membrane-coupled proteins. The apical plasma membrane is a central structure in mediating the function of the epithelium. It serves as an interface to produce and renew the apical extracellular matrix (aECM) that is essential for epithelial physiology and involves multiple epithelia-specific membranebound or membrane-associated effectors. These factors are not randomly distributed, but cluster with partners and occupy distinct subdomains within the membrane. Reflecting the non-random distribution of effectors, the apical plasma membrane adopts a typical topology that conceivably underlines its role in producing and influencing the extracellular milieu. The cues assembling subdomains, directing them to the correct site of function in the membrane, and possibly shaping the membrane are scarcely understood. Several epithelia in the Drosophila embryo provide a highly amenable model for a comparative investigation of apical plasma membrane modelling and equipment during development. In this review, we highlight these and summarise what is known about the underlying cellular and molecular processes.
Microvilli A common topological feature of the apical plasma membrane of many epithelia is an array of actin-stabilised microvilli, called the n
Corresponding author. E-mail address: [email protected] (B. Moussian).
0171-9335/$ - see front matter & 2009 Elsevier GmbH. All rights reserved. doi:10.1016/j.ejcb.2009.11.009
brush border (Danielsen and Hansen, 2003). The brush border provides an extensive apical membrane surface that assists the high absorptive and secretory activities of these epithelia. The microvilli subdivide the apical plasma membrane into two domains: the microvillus stalk and the invagination between two microvilli (Fig. 2A). The stalk carries enzymes and factors that are characteristic for the function of the epithelium, whereas the invaginations between the stalks constitute proposed sites of exoand endocytosis. The membrane-inserted mucin Muc1, for example, that contributes to the adhesive properties of certain epithelia, localises to the stalk of microvilli in cultured mammary epithelial cells (Bennett et al., 2001). The cytoskeletal support of the microvilli indicates that a microvillus is a static structure. Moreover, membrane proteins like Muc1 are connected to the microvillar actin filaments via actin-binding and -organising proteins such as ezrin (Bennett et al., 2001), agreeing with the notion of a fixed structure of the microvillus. In a recent work, however, McConnell and Tyska (2007) demonstrated that microvillus integrity may also involve dynamic processes mediated by myosin 1a that links the plasma membrane to the actin cytoskeleton. Myosin 1a in isolated rodent intestinal brush border drives the transport of membrane to the microvillar tips where it buds off and is released into the extracellular space as small vesicles (McConnell et al., 2009). Concomitantly, the microvillar stalk does not become shorter implying that for their full and continuous function microvilli have to be reconstituted, a process suggested to be driven by the fusion of Golgi-derived vesicles with the base of the microvilli. The factual impact of this process on the composition and function of the apical plasma membrane remains unclear. An unexpected notion about the dynamics of microvilli has been reported by Kultti et al. (2006). They have shown that the
ARTICLE IN PRESS A. Uv, B. Moussian / European Journal of Cell Biology 89 (2010) 208–211
Fig. 1. Epithelia are sheets of polarised cells with three types of plasma membrane surfaces. With the lateral plasma membrane epithelial cells communicate with their neighbours and establish paracellular barriers that separate a basal from an apical environment. At the basal surface, they usually are lined by an ECM, called the basal lamina that serves to contact other cell types or an interstitium. Their apical surface faces the external environment and often displays repetitive membrane protrusions that are involved to produce and organise an aECM. These protrusions are either insular (left) or longitudinal (right).
enzymatic activity of the transmembrane hyaluronic acid synthase HAS3 is sufficient to induce microvillus formation in cultured cells. This finding indicates that the apical plasma membrane is not simply a passive element responding to instructions by the cytoskeleton to shape a microvillus, but may actively contribute to its protrusion. In flies, microvilli are found in cuticle-free epithelia of the salivary gland, the Malpighian tubules and the midgut and have not been studied in detail. A glance at the brush border of the embryonic or larval midgut reveals even so structural and functional similarities between microvilli in mammals and Drosophila (Fig. 2A). Indeed, most factors associated with microvilli in mammalian cell culture are also found in Drosophila (Baumann, 2001; Edwards et al., 1997; Syed et al., 2008). For example, the midgut epithelium of the Drosophila embryo produces the peritrophic matrix that shares features with the vertebrate mucus. It consists of mucins and chitin that is synthesised by the membrane-inserted chitin synthase localising to the microvilli (Zimoch and Merzendorfer, 2002). Hence, the Drosophila embryo is an excellent model system to investigate microvilli formation and function in a multicellular and intact organism.
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cuticle is expanding uniformly axially and circumferentially, and that an axial restraint, possibly provided by the epithelium (but not the apical plasma membrane as it at first follows the buckles), causes the cuticle to buckle and form taenidial folds. The helical course of the taenidia could then be caused by some small randomly occurring torsional stresses in the tissue. Recent observations, however, point to an active role of the apical cell domain that includes the apical plasma membrane and the cortical cytoskeleton in taenidia formation. First, accumulation of electron-dense material only at the tips of the membrane folds, probably the chitin synthesis complex, indicates a non-uniform deposition of cuticle components, resulting in repetitive organisation of the taenidia. Second, Matusek et al. (2006) identified apical actin cables in the epithelium just prior to cuticle deposition, which define the future position of chitin synthesis forming the taenidial folds. Importantly, if the regularity of these actin rings or spirals is disrupted by the removal of an actin-organising formin, called DAAM (Dishevelled-associated activator of morphogenesis), the buckles still emerge but become correspondingly disorganised. It is possible that the subapical actin arrangement instructs the topology of the apical plasma membrane and causes clustering of specific biochemical functions to the apex of the membrane folds, and that this in turn determines the position of the taenidial folds. A fundamental problem is still the origin of taenidial fold/ subapical actin cytoskeleton geometry: What determines the orientation of the annular or helical rings surrounding the lumen, how do they continue undisrupted across cell boundaries, and what factors control the thickness and phasing of the taenidia? Are these properties solely controlled by the cytoskeleton or do they rely on functional interactions between the subapical actin organisation, the apical plasma membrane and aECM components? Is the apical plasma membrane topology and clustering of membrane activities the central link between epithelial cell organisation and the shape and properties of the aECM? We believe that the answers to these questions will be fundamental to our understanding of epithelial organ morphogenesis and physiology in general, and can be solved by mutational analyses in Drosophila.
The taenidia-forming apical plasma membrane of tracheal cells
The epidermal apical undulae
Two other types of apical membrane topologies in the developing fly embryo are found in the epidermis/hindgut and tracheal (respiratory) tubes, respectively. These epithelia synthesise and secrete the rigid protective cuticle, and during cuticle deposition their apical membrane exhibits repetitive folds that relate to the organisation of the cuticle. An essential feature of the tracheal cuticle is its arrangement into taenidial folds (Fig. 2B, C). The taenidia are ridges of procuticular material that assemble perpendicular to tube length, and form annular rings or spirals surrounding the lumen that run undisrupted across cell borders (Matusek et al., 2006). Deposition of the aECM components constituting the taenidial folds is preceded by the appearance of membrane folds that reflect the positions of the future taenidia (Fig. 2D). These membrane protrusions withdraw as the taenidial material is laid down, so that the apical plasma membrane becomes flattened beneath the growing ridges (Fig. 2F). The geometrical basis for the taenidial folds has aroused curiosity for decades. Thompson (1929) suggested that it ‘‘results from some simple physical laws and is produced by forces, which are unaffected by the existence of cell boundaries in the tracheal epithelium and act simply in the chitinous lining at the moment when it is being secreted’’. Locke (1958) hypothesised that the
Another type of regular protrusions of the apical plasma membrane, that we call apical undulae, is formed by epidermal and hindgut cells during cuticle production (Fig. 2E, H). These are longitudinal, microtubuli-underlain structures and run perpendicular to the anterior-posterior axis of the embryo (Moussian et al., 2006). They subdivide the apical plasma membrane into two domains, the crest carrying the chitin synthesis complex – the socalled plaque – and the valley separating two crests, where secretory vesicles fuse to release cuticular proteins into the extracellular space. The explicit configuration of the apical tier of the epidermal cells seemingly dictates the organisation of the extracellular matrix, as chitin fibres produced at the crests cross the apical undulae and are arranged parallel to the longitudinal axis of the embryo. Supporting this notion, mutations in syntaxin1A (syx1A) that abrogate apical undulae formation cause disorganisation of chitin fibres (Moussian et al., 2007). Since the secretion of the bulk of cuticle proteins also depends on the activity of the t-SNARE Syx1A, the disorganised chitin phenotype could arise from lack of interaction between chitin-organising proteins and chitin. Mutations in syx1A interfere with secretion and membrane trafficking without affecting partitioning of the apical plasma membrane into plaque and non-plaque domains. This observation
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Fig. 2. The apical cell domain of embryonic Drosophila epithelia. (A) Transmission electron microscopic (TEM) analysis of the Drosophila embryonic midgut reveals microvilli (mv) that cover the apical surface of the midgut epithelium. The microvilli are finger-like protrusions (arrow) stabilised by a rod of actin filaments. (B), (C) The taenidia (ta) of the tracheal cuticle run a helical course around the lumen and can be seen by light microscopy during late embryogenesis. With focus on the surface (B) they appear as dark lines perpendicular to tube length ((B), arrow), and when viewing longitudinal sections they are seen as dark punctae along the apical surface ((C), arrow). Stippled lines mark the basal epithelial surface. (D) TEM analysis of a longitudinal section of a tracheal tube reveals apical membrane (m) bulging during formation of the taenidial folds (e points to the envelope). Note the electron-dense areas at the tip of each membrane fold. (F) Apical membrane dynamics during taenidial formation: Initially, the membrane follows the outer lining of the cuticle (i), it then withdraws as taenidial aECM is deposited (ii) and later becomes flattened under the taenidial buckles (iii). ((E) and (H)) TEM analyses of the epithelial apical plasma membrane of the epidermis (E) and hindgut (H) show microtubuli-underlain longitudinal corrugations (apical undulae, u) during differentiation of the cuticle. Like the tracheal membrane folds, the undulae are electron-dense at the apex, but the cuticle surface (e) does not buckle accordingly. (G) Drawing of the epidermal apical undulae (u) in relation to the cuticular envelope (e). (I) The surface of the ventral epidermal cuticle exhibits rows of pigmented protrusions (left), called denticle belts (bracket), which are spaced by areas of naked cuticle. Scanning electron microscopic analyses illustrate the hook-like shape of the denticles (right). (J) Epidermal denticles or hairs disrupt the arrangement of the apical undulae. The plasma membrane at these sites carries a continuous plaque and forms actin-stabilised bulges to support the shape of the denticle or hair. (K) The chitin lamellar structure (bracket) in the procuticle as described by Yves Bouligand (1965) is disrupted at the denticle. The procuticle (pc) is marked in relation to the envelope (e) and the epicuticle (epc). Transmission electron micrographs were taken on a Philips CM10 electron microscope at 60 kV from samples prepared as described by Moussian et al. (2006).
suggests that the separation of membrane domains requires Syx1A-independent mechanisms. Syx1A is also dispensable for the assembly of the plaques, indicating that at least one other t-SNARE is needed for a complete equipment of the epidermal apical plasma membrane. Moreover, Syx1A is present in the epidermal cells prior to cuticle deposition. Consequently, its cellular role has to be switched from a non-undular to an undula-forming function. Taken together, correct cuticle differentiation necessitates temporal and spatial coordination of several membrane-modelling and -decorating mechanisms. The factors participating at these fascinating processes are to date totally obscure.
The apical undulae do not homogenously occupy the apical surface of epidermal cells. At positions where epidermal hairs and denticles are formed, the apical plasma membrane bulges, disrupting the regularity of the apical undulae (Fig. 2I, J). These bulges engage vertical bundles of actin fibres and microtubules, as well as specialised membrane proteins such as Miniature and Dusky (Chanut-Delalande et al., 2006), members of the zonula pellucida (ZP) class of proteins. Conceptually, membrane zones defined by ZP proteins could suppress the mechanisms of apical undulae establishment by preventing the reorganisation of the cytoskeleton parallel to the plasma membrane. Consistent with this view, membrane bulges leading to hair or denticle formation
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emerge before the apical undulae are sculpted. Probably due to the lack of apical undulae, chitin fibres at the positions of hair or denticle membrane bulging are not organised as in the cuticle deposited at sites harbouring normal apical undulae (Fig. 2K). Despite their similar apical plasma membrane topology, tracheal and epidermal cells configure different types of cuticle: the taenidial cuticle obeys the corrugated shape of the apical plasma membrane, while the epidermal cuticle adopts a uniform thickness (Fig. 2D, E). This fundamental difference implies tissuespecific implementation and regulation of membrane-inherent as well as extracellular activities. The factors that account for these processes await identification.
to its own needs. The molecular determinants of the apical plasma membrane interacting either with the aECM or the cytoskeleton have only been poorly investigated. The Drosophila embryo harbours a number of differently shaped apical plasma membranes that play a key role during terminal differentiation of organs. Investigation of the molecular mechanisms modelling and equipping the apical plasma membrane in these epithelial tissues of Drosophila will enhance our understanding of cell differentiation.
The apical plasma membrane at the onset of cuticle production
B. Moussian is supported by the German Research Foundation (DFG MO1714 and SFB 446/A29 ‘‘Mechanisms of Cell Behaviour in Eukaryotes’’). A. Uv is supported by the Swedish Research Council (VR). We thank the Wenner-Gren Foundation for financial support. We are grateful to Kate Kearney for comments on the manuscript.
Prior to formation of the apical undulae the apical plasma membrane protrudes at those sites where the first and outermost cuticle layer – the envelope – is being assembled. In contrast to the microvilli, the apical undulae and the taenidiae, these protrusions occur irregularly. The random allocation of envelope-depositing sites suggests spontaneous and non-patterned recruitment of cytoplasmic components, including the cytoskeleton, by the plasma membrane. In other words, membraneinserted and -associated proteins and enzymes producing the envelope and ejecting it into the extracellular space may induce membrane bending and bulging. This scenario is in line with the above-mentioned work demonstrating that the enzymatic activity of the transmembrane HAS3 in cultured epithelial cells provokes formation of microvilli-like membrane protrusions (Kultti et al., 2006). In both instances, the underlying forces may originate from the extracellular matrix (envelope or hyaluronic acid) or from the membrane itself. A detailed study of the factors and mechanisms responsible for this phenomenon is missing. After envelope formation, the apical plasma membrane switches from a random activity to a patterned one during epiand procuticle assembly. This switch is accompanied by massive secretion of cuticle material and chitin synthesis implying considerable reengineering of secretory and membrane trafficking routes and probably the rearrangement of the subapical cytoskeleton. The signals and mechanisms adjusting the function of the apical plasma membrane during the different steps of cuticle differentiation are unknown. The exploration of the players involved is certainly of broad interest, as transitions of plasma membrane activity are likely to be common events in epithelia that produce an aECM.
Conclusion At the apical domain of epithelial cells there is an intimate relationship between the aECM, the apical plasma membrane and the cytosolic subapical cytoskeleton. The specific topology of the membrane and, consequently, the asymmetric distribution of membrane-bound and -associated effectors obviously depend on the underlain cytoskeleton, in turn influencing the organisation of the aECM. Several observations suggest that the apical plasma membrane does not simply obey instructions of the cytoskeleton, but may have the ability to reorganise the cytoskeleton according
Acknowledgements
References Baumann, O., 2001. Posterior midgut epithelial cells differ in their organization of the membrane skeleton from other Drosophila epithelia. Exp. Cell Res. 270, 176–187. Bennett Jr., R., Jarvela, T., Engelhardt, P., Kostamovaara, L., Sparks, P., Carpen, O., Turunen, O., Vaheri, A., 2001. Mucin MUC1 is seen in cell surface protrusions together with ezrin in immunoelectron tomography and is concentrated at tips of filopodial protrusions in MCF-7 breast carcinoma cells. J. Histochem. Cytochem. 49, 67–77. Bouligand, Y., 1965. On a twisted fibrillar arrangement common to several biologic structures. CR Acad. Sci. Hebd. Seances Acad. Sci. D 261, 4864–4867 [French]. Chanut-Delalande, H., Fernandes, I., Roch, F., Payre, F., Plaza, S., 2006. Shavenbaby couples patterning to epidermal cell shape control. PLoS Biol. 4, e290. Danielsen, E.M., Hansen, G.H., 2003. Lipid rafts in epithelial brush borders: atypical membrane microdomains with specialized functions. Biochim. Biophys. Acta 1617, 1–9. Edwards, K.A., Demsky, M., Montague, R.A., Weymouth, N., Kiehart, D.P., 1997. GFP-moesin illuminates actin cytoskeleton dynamics in living tissue and demonstrates cell shape changes during morphogenesis in Drosophila. Dev. Biol. 191, 103–117. Kultti, A., Rilla, K., Tiihonen, R., Spicer, A.P., Tammi, R.H., Tammi, M.I., 2006. Hyaluronan synthesis induces microvillus-like cell surface protrusions. J. Biol. Chem. 281, 15821–15828. Locke, M., 1958. The formation of tracheae and tracheoles in Rhodnius prolixus. Q. J. Microsc. Sci. s3-99, 29–46. Matusek, T., Djiane, A., Jankovics, F., Brunner, D., Mlodzik, M., Mihaly, J., 2006. The Drosophila formin DAAM regulates the tracheal cuticle pattern through organizing the actin cytoskeleton. Development 133, 957–966. McConnell, R.E., Tyska, M.J., 2007. Myosin-1a powers the sliding of apical membrane along microvillar actin bundles. J. Cell Biol 177, 671–681. McConnell, R.E., Higginbotham, J.N., Shifrin Jr., D.A., Tabb, D.L., Coffey, R.J., Tyska, M.J., 2009. The enterocyte microvillus is a vesicle-generating organelle. J. Cell Biol. 185, 1285–1298. Moussian, B., Seifarth, C., Muller, U., Berger, J., Schwarz, H., 2006. Cuticle differentiation during Drosophila embryogenesis. Arthropod Struct. Dev. 35, 137–152. Moussian, B., Veerkamp, J., Muller, U., Schwarz, H., 2007. Assembly of the Drosophila larval exoskeleton requires controlled secretion and shaping of the apical plasma membrane. Matrix Biol. 26, 337–347. Syed, Z.A., Hard, T., Uv, A., van Dijk-Hard, I.F., 2008. A potential role for Drosophila mucins in development and physiology. PLoS One 3, e3041. Tepass, U., Hartenstein, V., 1994. The development of cellular junctions in the Drosophila embryo. Dev. Biol. 161, 563–596. Thompson, W.R., 1929. A contribution to the study of the morphogenesis in the muscoid Diptera. Trans. Roy. Ent. Soc. Lond. 77, 195–244. Zimoch, L., Merzendorfer, H., 2002. Immunolocalization of chitin synthase in the tobacco hornworm. Cell Tissue Res. 308, 287–297.
Contents lists available at ScienceDirect
European Journal of Cell Biology journal homepage: www.elsevier.de/ejcb
The apical plasma membrane of Drosophila embryonic epithelia Anne Uv a, Bernard Moussian b,n a b
Institute of Biomedicine, Department of Medicinal Genetics, Gothenburg University, Medicinaregatan 9A, 40530 Gothenburg, Sweden Max-Planck Institute for Developmental Biology, Spemannstr. 35, 72076 T¨ ubingen, Germany
a r t i c l e in f o
Keywords: Epithelia Epidermis Trachea Midgut Apical plasma membrane Microvilli Cuticle Taenidia Apical undulae
a b s t r a c t The apical plasma membrane of epithelia presents the interface between organs and the external environment. It has biochemical activities distinct from those of the basal and lateral plasma membranes, as it accommodates the production and assembly of ordered apical matrices involved in organ protection and physiology and determines the microenvironment in the apical extracellular milieu. Here, we emphasise the importance of the apical plasma membrane in tissue differentiation, by mainly focussing on the embryo of the fruit fly Drosophila melanogaster, and discuss the principal organisation of the apical plasma membrane into repetitive subdomains of specific topologies and activities essential for epithelial function. & 2009 Elsevier GmbH. All rights reserved.
Introduction The plasma membrane of epithelial cells is subdivided into three functional areas, the apical, the lateral and the basal plasma membrane (Fig. 1) (Tepass and Hartenstein, 1994), that are defined by distinct membrane-coupled proteins. The apical plasma membrane is a central structure in mediating the function of the epithelium. It serves as an interface to produce and renew the apical extracellular matrix (aECM) that is essential for epithelial physiology and involves multiple epithelia-specific membranebound or membrane-associated effectors. These factors are not randomly distributed, but cluster with partners and occupy distinct subdomains within the membrane. Reflecting the non-random distribution of effectors, the apical plasma membrane adopts a typical topology that conceivably underlines its role in producing and influencing the extracellular milieu. The cues assembling subdomains, directing them to the correct site of function in the membrane, and possibly shaping the membrane are scarcely understood. Several epithelia in the Drosophila embryo provide a highly amenable model for a comparative investigation of apical plasma membrane modelling and equipment during development. In this review, we highlight these and summarise what is known about the underlying cellular and molecular processes.
Microvilli A common topological feature of the apical plasma membrane of many epithelia is an array of actin-stabilised microvilli, called the n
Corresponding author. E-mail address: [email protected] (B. Moussian).
0171-9335/$ - see front matter & 2009 Elsevier GmbH. All rights reserved. doi:10.1016/j.ejcb.2009.11.009
brush border (Danielsen and Hansen, 2003). The brush border provides an extensive apical membrane surface that assists the high absorptive and secretory activities of these epithelia. The microvilli subdivide the apical plasma membrane into two domains: the microvillus stalk and the invagination between two microvilli (Fig. 2A). The stalk carries enzymes and factors that are characteristic for the function of the epithelium, whereas the invaginations between the stalks constitute proposed sites of exoand endocytosis. The membrane-inserted mucin Muc1, for example, that contributes to the adhesive properties of certain epithelia, localises to the stalk of microvilli in cultured mammary epithelial cells (Bennett et al., 2001). The cytoskeletal support of the microvilli indicates that a microvillus is a static structure. Moreover, membrane proteins like Muc1 are connected to the microvillar actin filaments via actin-binding and -organising proteins such as ezrin (Bennett et al., 2001), agreeing with the notion of a fixed structure of the microvillus. In a recent work, however, McConnell and Tyska (2007) demonstrated that microvillus integrity may also involve dynamic processes mediated by myosin 1a that links the plasma membrane to the actin cytoskeleton. Myosin 1a in isolated rodent intestinal brush border drives the transport of membrane to the microvillar tips where it buds off and is released into the extracellular space as small vesicles (McConnell et al., 2009). Concomitantly, the microvillar stalk does not become shorter implying that for their full and continuous function microvilli have to be reconstituted, a process suggested to be driven by the fusion of Golgi-derived vesicles with the base of the microvilli. The factual impact of this process on the composition and function of the apical plasma membrane remains unclear. An unexpected notion about the dynamics of microvilli has been reported by Kultti et al. (2006). They have shown that the
ARTICLE IN PRESS A. Uv, B. Moussian / European Journal of Cell Biology 89 (2010) 208–211
Fig. 1. Epithelia are sheets of polarised cells with three types of plasma membrane surfaces. With the lateral plasma membrane epithelial cells communicate with their neighbours and establish paracellular barriers that separate a basal from an apical environment. At the basal surface, they usually are lined by an ECM, called the basal lamina that serves to contact other cell types or an interstitium. Their apical surface faces the external environment and often displays repetitive membrane protrusions that are involved to produce and organise an aECM. These protrusions are either insular (left) or longitudinal (right).
enzymatic activity of the transmembrane hyaluronic acid synthase HAS3 is sufficient to induce microvillus formation in cultured cells. This finding indicates that the apical plasma membrane is not simply a passive element responding to instructions by the cytoskeleton to shape a microvillus, but may actively contribute to its protrusion. In flies, microvilli are found in cuticle-free epithelia of the salivary gland, the Malpighian tubules and the midgut and have not been studied in detail. A glance at the brush border of the embryonic or larval midgut reveals even so structural and functional similarities between microvilli in mammals and Drosophila (Fig. 2A). Indeed, most factors associated with microvilli in mammalian cell culture are also found in Drosophila (Baumann, 2001; Edwards et al., 1997; Syed et al., 2008). For example, the midgut epithelium of the Drosophila embryo produces the peritrophic matrix that shares features with the vertebrate mucus. It consists of mucins and chitin that is synthesised by the membrane-inserted chitin synthase localising to the microvilli (Zimoch and Merzendorfer, 2002). Hence, the Drosophila embryo is an excellent model system to investigate microvilli formation and function in a multicellular and intact organism.
209
cuticle is expanding uniformly axially and circumferentially, and that an axial restraint, possibly provided by the epithelium (but not the apical plasma membrane as it at first follows the buckles), causes the cuticle to buckle and form taenidial folds. The helical course of the taenidia could then be caused by some small randomly occurring torsional stresses in the tissue. Recent observations, however, point to an active role of the apical cell domain that includes the apical plasma membrane and the cortical cytoskeleton in taenidia formation. First, accumulation of electron-dense material only at the tips of the membrane folds, probably the chitin synthesis complex, indicates a non-uniform deposition of cuticle components, resulting in repetitive organisation of the taenidia. Second, Matusek et al. (2006) identified apical actin cables in the epithelium just prior to cuticle deposition, which define the future position of chitin synthesis forming the taenidial folds. Importantly, if the regularity of these actin rings or spirals is disrupted by the removal of an actin-organising formin, called DAAM (Dishevelled-associated activator of morphogenesis), the buckles still emerge but become correspondingly disorganised. It is possible that the subapical actin arrangement instructs the topology of the apical plasma membrane and causes clustering of specific biochemical functions to the apex of the membrane folds, and that this in turn determines the position of the taenidial folds. A fundamental problem is still the origin of taenidial fold/ subapical actin cytoskeleton geometry: What determines the orientation of the annular or helical rings surrounding the lumen, how do they continue undisrupted across cell boundaries, and what factors control the thickness and phasing of the taenidia? Are these properties solely controlled by the cytoskeleton or do they rely on functional interactions between the subapical actin organisation, the apical plasma membrane and aECM components? Is the apical plasma membrane topology and clustering of membrane activities the central link between epithelial cell organisation and the shape and properties of the aECM? We believe that the answers to these questions will be fundamental to our understanding of epithelial organ morphogenesis and physiology in general, and can be solved by mutational analyses in Drosophila.
The taenidia-forming apical plasma membrane of tracheal cells
The epidermal apical undulae
Two other types of apical membrane topologies in the developing fly embryo are found in the epidermis/hindgut and tracheal (respiratory) tubes, respectively. These epithelia synthesise and secrete the rigid protective cuticle, and during cuticle deposition their apical membrane exhibits repetitive folds that relate to the organisation of the cuticle. An essential feature of the tracheal cuticle is its arrangement into taenidial folds (Fig. 2B, C). The taenidia are ridges of procuticular material that assemble perpendicular to tube length, and form annular rings or spirals surrounding the lumen that run undisrupted across cell borders (Matusek et al., 2006). Deposition of the aECM components constituting the taenidial folds is preceded by the appearance of membrane folds that reflect the positions of the future taenidia (Fig. 2D). These membrane protrusions withdraw as the taenidial material is laid down, so that the apical plasma membrane becomes flattened beneath the growing ridges (Fig. 2F). The geometrical basis for the taenidial folds has aroused curiosity for decades. Thompson (1929) suggested that it ‘‘results from some simple physical laws and is produced by forces, which are unaffected by the existence of cell boundaries in the tracheal epithelium and act simply in the chitinous lining at the moment when it is being secreted’’. Locke (1958) hypothesised that the
Another type of regular protrusions of the apical plasma membrane, that we call apical undulae, is formed by epidermal and hindgut cells during cuticle production (Fig. 2E, H). These are longitudinal, microtubuli-underlain structures and run perpendicular to the anterior-posterior axis of the embryo (Moussian et al., 2006). They subdivide the apical plasma membrane into two domains, the crest carrying the chitin synthesis complex – the socalled plaque – and the valley separating two crests, where secretory vesicles fuse to release cuticular proteins into the extracellular space. The explicit configuration of the apical tier of the epidermal cells seemingly dictates the organisation of the extracellular matrix, as chitin fibres produced at the crests cross the apical undulae and are arranged parallel to the longitudinal axis of the embryo. Supporting this notion, mutations in syntaxin1A (syx1A) that abrogate apical undulae formation cause disorganisation of chitin fibres (Moussian et al., 2007). Since the secretion of the bulk of cuticle proteins also depends on the activity of the t-SNARE Syx1A, the disorganised chitin phenotype could arise from lack of interaction between chitin-organising proteins and chitin. Mutations in syx1A interfere with secretion and membrane trafficking without affecting partitioning of the apical plasma membrane into plaque and non-plaque domains. This observation
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Fig. 2. The apical cell domain of embryonic Drosophila epithelia. (A) Transmission electron microscopic (TEM) analysis of the Drosophila embryonic midgut reveals microvilli (mv) that cover the apical surface of the midgut epithelium. The microvilli are finger-like protrusions (arrow) stabilised by a rod of actin filaments. (B), (C) The taenidia (ta) of the tracheal cuticle run a helical course around the lumen and can be seen by light microscopy during late embryogenesis. With focus on the surface (B) they appear as dark lines perpendicular to tube length ((B), arrow), and when viewing longitudinal sections they are seen as dark punctae along the apical surface ((C), arrow). Stippled lines mark the basal epithelial surface. (D) TEM analysis of a longitudinal section of a tracheal tube reveals apical membrane (m) bulging during formation of the taenidial folds (e points to the envelope). Note the electron-dense areas at the tip of each membrane fold. (F) Apical membrane dynamics during taenidial formation: Initially, the membrane follows the outer lining of the cuticle (i), it then withdraws as taenidial aECM is deposited (ii) and later becomes flattened under the taenidial buckles (iii). ((E) and (H)) TEM analyses of the epithelial apical plasma membrane of the epidermis (E) and hindgut (H) show microtubuli-underlain longitudinal corrugations (apical undulae, u) during differentiation of the cuticle. Like the tracheal membrane folds, the undulae are electron-dense at the apex, but the cuticle surface (e) does not buckle accordingly. (G) Drawing of the epidermal apical undulae (u) in relation to the cuticular envelope (e). (I) The surface of the ventral epidermal cuticle exhibits rows of pigmented protrusions (left), called denticle belts (bracket), which are spaced by areas of naked cuticle. Scanning electron microscopic analyses illustrate the hook-like shape of the denticles (right). (J) Epidermal denticles or hairs disrupt the arrangement of the apical undulae. The plasma membrane at these sites carries a continuous plaque and forms actin-stabilised bulges to support the shape of the denticle or hair. (K) The chitin lamellar structure (bracket) in the procuticle as described by Yves Bouligand (1965) is disrupted at the denticle. The procuticle (pc) is marked in relation to the envelope (e) and the epicuticle (epc). Transmission electron micrographs were taken on a Philips CM10 electron microscope at 60 kV from samples prepared as described by Moussian et al. (2006).
suggests that the separation of membrane domains requires Syx1A-independent mechanisms. Syx1A is also dispensable for the assembly of the plaques, indicating that at least one other t-SNARE is needed for a complete equipment of the epidermal apical plasma membrane. Moreover, Syx1A is present in the epidermal cells prior to cuticle deposition. Consequently, its cellular role has to be switched from a non-undular to an undula-forming function. Taken together, correct cuticle differentiation necessitates temporal and spatial coordination of several membrane-modelling and -decorating mechanisms. The factors participating at these fascinating processes are to date totally obscure.
The apical undulae do not homogenously occupy the apical surface of epidermal cells. At positions where epidermal hairs and denticles are formed, the apical plasma membrane bulges, disrupting the regularity of the apical undulae (Fig. 2I, J). These bulges engage vertical bundles of actin fibres and microtubules, as well as specialised membrane proteins such as Miniature and Dusky (Chanut-Delalande et al., 2006), members of the zonula pellucida (ZP) class of proteins. Conceptually, membrane zones defined by ZP proteins could suppress the mechanisms of apical undulae establishment by preventing the reorganisation of the cytoskeleton parallel to the plasma membrane. Consistent with this view, membrane bulges leading to hair or denticle formation
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emerge before the apical undulae are sculpted. Probably due to the lack of apical undulae, chitin fibres at the positions of hair or denticle membrane bulging are not organised as in the cuticle deposited at sites harbouring normal apical undulae (Fig. 2K). Despite their similar apical plasma membrane topology, tracheal and epidermal cells configure different types of cuticle: the taenidial cuticle obeys the corrugated shape of the apical plasma membrane, while the epidermal cuticle adopts a uniform thickness (Fig. 2D, E). This fundamental difference implies tissuespecific implementation and regulation of membrane-inherent as well as extracellular activities. The factors that account for these processes await identification.
to its own needs. The molecular determinants of the apical plasma membrane interacting either with the aECM or the cytoskeleton have only been poorly investigated. The Drosophila embryo harbours a number of differently shaped apical plasma membranes that play a key role during terminal differentiation of organs. Investigation of the molecular mechanisms modelling and equipping the apical plasma membrane in these epithelial tissues of Drosophila will enhance our understanding of cell differentiation.
The apical plasma membrane at the onset of cuticle production
B. Moussian is supported by the German Research Foundation (DFG MO1714 and SFB 446/A29 ‘‘Mechanisms of Cell Behaviour in Eukaryotes’’). A. Uv is supported by the Swedish Research Council (VR). We thank the Wenner-Gren Foundation for financial support. We are grateful to Kate Kearney for comments on the manuscript.
Prior to formation of the apical undulae the apical plasma membrane protrudes at those sites where the first and outermost cuticle layer – the envelope – is being assembled. In contrast to the microvilli, the apical undulae and the taenidiae, these protrusions occur irregularly. The random allocation of envelope-depositing sites suggests spontaneous and non-patterned recruitment of cytoplasmic components, including the cytoskeleton, by the plasma membrane. In other words, membraneinserted and -associated proteins and enzymes producing the envelope and ejecting it into the extracellular space may induce membrane bending and bulging. This scenario is in line with the above-mentioned work demonstrating that the enzymatic activity of the transmembrane HAS3 in cultured epithelial cells provokes formation of microvilli-like membrane protrusions (Kultti et al., 2006). In both instances, the underlying forces may originate from the extracellular matrix (envelope or hyaluronic acid) or from the membrane itself. A detailed study of the factors and mechanisms responsible for this phenomenon is missing. After envelope formation, the apical plasma membrane switches from a random activity to a patterned one during epiand procuticle assembly. This switch is accompanied by massive secretion of cuticle material and chitin synthesis implying considerable reengineering of secretory and membrane trafficking routes and probably the rearrangement of the subapical cytoskeleton. The signals and mechanisms adjusting the function of the apical plasma membrane during the different steps of cuticle differentiation are unknown. The exploration of the players involved is certainly of broad interest, as transitions of plasma membrane activity are likely to be common events in epithelia that produce an aECM.
Conclusion At the apical domain of epithelial cells there is an intimate relationship between the aECM, the apical plasma membrane and the cytosolic subapical cytoskeleton. The specific topology of the membrane and, consequently, the asymmetric distribution of membrane-bound and -associated effectors obviously depend on the underlain cytoskeleton, in turn influencing the organisation of the aECM. Several observations suggest that the apical plasma membrane does not simply obey instructions of the cytoskeleton, but may have the ability to reorganise the cytoskeleton according
Acknowledgements
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