Adherens Junction: Molecular Architecture and Regulation

The adherens junction (AJ) is an element of the cell–cell junction in which cadherin receptors bridge the neighboring plasma membranes via their homophilic interactions. Cadherins associate with cytoplasmic proteins, called catenins, which in turn bind to cytoskeletal components, such as actin filaments and microtubules. These molecular complexes further interact with other proteins, including signaling molecules, rendering the AJs into highly dynamic and regulatable structures. The AJs of such nature contribute to the physical linking of cells, as well as to the regulation of cell–cell contacts, which is essential for morphogenesis and remodeling of tissues and organs. Thus, elucidating the molecular architecture of the AJs and their regulatory mechanisms are crucial for understanding how the multicellular system is organized.The adherens junction (AJ) is a form of cell–cell adhesion structure observed in a variety of cell types, as well as in different animal species. It is characterized by a pair of plasma membranes apposed with a distance of 10–20 nm between them, whose intercellular space is occupied by rod-shaped molecules bridging the membranes (Hirokawa and Heuser 1981; Miyaguchi 2000), and the cytoplasmic side of the AJ is associated with condensed actin filaments. In polarized epithelia of vertebrates, the AJ is part of the tripartite junctional complex localized at the juxta-luminal region, which comprises the tight junction (zonula occludens), AJ, and desmosome (macula adherens) aligned in this order from the apical end of the junction (Farquhar and Palade 1963). In this type of epithelia, the AJ is specifically termed the “zonula adherens” or “adhesion belt,” as it completely encloses the cells along with the F-actin lining, called the circumferential actin belt (Fig. 1). The AJs in other cell types assume different morphologies: For example, the AJs in fibroblastic cells are spotty and discontinuous (Yonemura et al. 1995), and those in neurons are organized into tiny puncta as a constituent of the synaptic junctions (Uchida et al. 1996).Open in a separate windowFigure 1.Morphological variations of the adherens junction. In Caco2 cells (colonic carcinoma line), E-cadherin is localized along the actin circumferential belt to organize the zonula adherens (arrow). At the lateral portions of cell junction (arrowheads), E-cadherin signals are punctate, only occasionally overlapping with actin signals in this specific sample. The lateral patterns of cadherin and actin distribution, however, vary with cellular conditions. In MCF10A cells (mammary epithelial line), spotty adherens junctions are seen, where actin filaments perpendicularly terminate at E-cadherin puncta.A major function of AJs is to maintain the physical association between cells, as disruption of them causes loosening of cell–cell contacts, leading to disorganization of tissue architecture. Calcium chelators such as EDTA and EGTA are widely used as a reagent to promote the dissociation of cells in tissues or monolayer cultures. A major target of these chelators is the AJ, as this is a calcium-sensitive structure; although, calcium removal is generally insufficient for the complete dispersion of cells because of the presence of calcium-independent cell–cell adhesion mechanisms (Takeichi et al. 1977). Early studies to search for the molecules responsible for the calcium-dependent junctions resulted in the identification of a group of type-I transmembrane proteins, and its founding member was termed cadherin (Yoshida and Takeichi 1982; Yoshida-Noro et al. 1984). Related molecules identified were also called by various names, such as uvomorulin (Peyrieras et al. 1983), LCAM (Gallin et al. 1983), and ACAM (Volk and Geiger 1984). Later studies revealed that the cadherins form a superfamily, and therefore, the original cadherins are now called “classic” cadherins.Another series of studies have identified nectins, a family of immunoglobulin-like transmembrane proteins, as an AJ component. Nectins function in a calcium-independent way to promote cell–cell adhesion (Nakanishi and Takai 2004). In this article, we overview the molecular organization of the AJs constructed with these membrane proteins, as well as the regulatory mechanisms that operate to sustain or remodel these junctions, paying much attention to the linkages between the AJ and cytoskeletal or signaling proteins.     

Molecular Bases of Cell–Cell Junctions Stability and Dynamics

Epithelial cell–cell junctions are formed by apical adherens junctions (AJs), which are composed of cadherin adhesion molecules interacting in a dynamic way with the cortical actin cytoskeleton. Regulation of cell–cell junction stability and dynamics is crucial to maintain tissue integrity and allow tissue remodeling throughout development. Actin filament turnover and organization are tightly controlled together with myosin-II activity to produce mechanical forces that drive the assembly, maintenance, and remodeling of AJs. In this review, we will discuss these three distinct stages in the lifespan of cell–cell junctions, using several developmental contexts, which illustrate how mechanical forces are generated and transmitted at junctions, and how they impact on the integrity and the remodeling of cell–cell junctions.Cell–cell junction formation and remodeling occur repeatedly throughout development. Epithelial cells are linked by apical adherens junctions (AJs) that rely on the cadherin-catenin-actin module. Cadherins, of which epithelial E-cadherin (E-cad) is the most studied, are Ca²⁺-dependent transmembrane adhesion proteins forming homophilic and heterophilic bonds in trans between adjacent cells. Cadherins and the actin cytoskeleton are mutually interdependent (Jaffe et al. 1990; Matsuzaki et al. 1990; Hirano et al. 1992; Oyama et al. 1994; Angres et al. 1996; Orsulic and Peifer 1996; Adams et al. 1998; Zhang et al. 2005; Pilot et al. 2006). This has long been attributed to direct physical interaction of E-cad with β-catenin (β-cat) and of α-catenin (α-cat) with actin filaments (for reviews, see Gumbiner 2005; Leckband and Prakasam 2006; Pokutta and Weis 2007). Recently, biochemical and protein dynamics analyses have shown that such a link may not exist and that instead, a constant shuttling of α-cat between cadherin/β-cat complexes and actin may be key to explain the dynamic aspect of cell–cell adhesion (Drees et al. 2005; Yamada et al. 2005). Regardless of the exact nature of this link, several studies show that AJs are indeed physically attached to actin and that cadherins transmit cortical forces exerted by junctional acto-myosin networks (Costa et al. 1998; Sako et al. 1998; Pettitt et al. 2003; Dawes-Hoang et al. 2005; Cavey et al. 2008; Martin et al. 2008; Rauzi et al. 2008). In addition, physical association depends in part on α-cat (Cavey et al. 2008) and additional intermediates have been proposed to represent alternative missing links (Abe and Takeichi 2008) (reviewed in Gates and Peifer 2005; Weis and Nelson 2006). Although further work is needed to address the molecular nature of cadherin/actin dynamic interactions, association with actin is crucial all throughout the lifespan of AJs. In this article, we will review our current understanding of the molecular mechanisms at work during three different developmental stages of AJs biology: assembly, stabilization, and remodeling, with special emphasis on the mechanical forces controlling AJs integrity and development.     

Presynaptic Membrane Retrieval and Endosome Biology: Defining Molecularly Heterogeneous Synaptic Vesicles

The release and uptake of neurotransmitters by synaptic vesicles is a tightly controlled process that occurs in response to diverse stimuli at morphologically disparate synapses. To meet these architectural and functional synaptic demands, it follows that there should be diversity in the mechanisms that control their secretion and retrieval and possibly in the composition of synaptic vesicles within the same terminal. Here we pay particular attention to areas where such diversity is generated, such as the variance in exocytosis/endocytosis coupling, SNAREs defining functionally diverse synaptic vesicle populations and the adaptor-dependent sorting machineries capable of generating vesicle diversity. We argue that there are various synaptic vesicle recycling pathways at any given synapse and discuss several lines of evidence that support the role of the endosome in synaptic vesicle recycling.Chemical synapses contain discrete numbers of synaptic vesicles, which are capable of sustaining neurotransmitter release. Sustained neurotransmission occurs despite the secretory demands imposed by persistent and diverse patterns of neuronal electrical activity. Maintaining synaptic vesicle numbers requires local mechanisms to regenerate these vesicles to prevent their exhaustion, preserve plasma membrane surface area, and to maintain the molecularly distinct identity of a vesicle versus plasma membrane. Rizzoli and Betz (2005) eloquently draw a parallel between chemical neurotransmission with synapse chatter saying that some synapses “whisper,” whereas others “shout.” The “louder” the synapse, the more synaptic vesicles are required, extending from a few hundred vesicles (whisperers) to nearly thousands (shouters). This beautiful analogy implies that every synapse has just one “voice” or species of vesicle. Here we will present the case that synapses are more like choirs in which multiple vesicle species or “voices” contribute to the “pianissimo” or “fortissimo” parts of chemical neurotransmission.Synaptic terminals show a range of structural and functional differences in distinct regions of the brain, suggesting that the mechanisms for exocytosis/endocytosis coupling, as well as local vesicle recycling, may also be diverse. On one side, the Calyx of Held nerve terminal participates in fast and sustained synaptic transmission at high frequency (800 Hz), which is crucial for sound localization in the auditory brainstem (Taschenberger and von Gersdorff 2000; Borst and Soria van Hoeve 2012). The Calyx of Held houses ∼70,000 synaptic vesicles with nearly 3000 vesicles docked per Calyx terminal. These docked vesicles are distributed across the ∼500 active zones that exist per Calyx where vesicle fusion occurs (Satzler et al. 2002). On the other hand, hippocampal synapses fire action potentials at ∼0.5 Hz in bursts (Dobrunz and Stevens 1999). This synapse contains ∼200 synaptic vesicles and one active zone with ∼10 vesicles docked (Schikorski and Stevens 1997). With such a wide functional and structural gamut of synapses, it is reasonable to hypothesize that synaptic vesicles may differ in their retrieval mechanisms, not just at the rate at which the process occurs but also in the molecular pathways used.Two synaptic vesicle retrieval mechanisms, namely clathrin/AP-2/dynamin-dependent biogenesis and kiss-and-run, have been summarized in outstanding recent reviews (see, for example, Augustine et al. 2006; Rizzoli and Jahn 2007; Smith et al. 2008; Royle and Lagnado 2010; Ferguson and De Camilli 2012; Saheki and De Camilli 2012). Therefore, here we focus on the coupling of secretion and membrane retrieval, as well as endosome sorting. We will discuss new developments supporting the existence of diverse functional and molecular pools of synaptic vesicles and how endocytosis and endosome retrieval mechanisms may generate these vesicle pools.     

The Desmosome

Desmosomes are intercellular junctions that tether intermediate filaments to the plasma membrane. Desmogleins and desmocollins, members of the cadherin superfamily, mediate adhesion at desmosomes. Cytoplasmic components of the desmosome associate with the desmosomal cadherin tails through a series of protein interactions, which serve to recruit intermediate filaments to sites of desmosome assembly. These desmosomal plaque components include plakoglobin and the plakophilins, members of the armadillo gene family. Linkage to the cytoskeleton is mediated by the intermediate filament binding protein, desmoplakin, which associates with both plakoglobin and plakophilins. Although desmosomes are critical for maintaining stable cell–cell adhesion, emerging evidence indicates that they are also dynamic structures that contribute to cellular processes beyond that of cell adhesion. This article outlines the structure and function of the major desmosomal proteins, and explores the contributions of this protein complex to tissue architecture and morphogenesis.The desmosome is an adhesive intercellular junction that is crucial to tissues that experience mechanical stress, such as the myocardium, bladder, gastrointestinal mucosa, and skin (Getsios et al. 2004b; Holthofer et al. 2007). The desmosome was first observed in the spinous layer of epidermis by the Italian pathologist Giulio Bizzozero (1846–1901). Bizzozero''s observations of these small dense nodules, subsequently named “nodes of Bizzozero,” led him to the insightful interpretation of these structures as adhesive cell–cell contact points. The term desmosome was later coined by Josef Schaffer in 1920 and is derived from the Greek words “desmo,” meaning bond or fastening, and “soma,” meaning body (Wells 2005; Calkins and Setzer 2007). The introduction of electron microscopy yielded a series of advances by Porter, Odland, and Kelly in the 1950s and 1960s, which revealed desmosome organization at the ultrastructural level. These studies and others indicated that the desmosome can be divided into three morphologically identifiable zones: the extracellular core region (desmoglea), the outer dense plaque (ODP), and the inner dense plaque (IDP) (Fig. 1A) (Kowalczyk et al. 1994; Schmidt et al. 1994; Green and Jones 1996; North et al. 1999; Garrod and Chidgey 2008).Open in a separate windowFigure 1.A model for the structure of desmosomes. (A) Electron micrograph of a desmosome. (B) Schematic of desmosomal proteins and relative distance from the plasma membrane (PM). The desmosomal cadherins, the desmogleins and desmocollins, extend into extracellular core and outer dense plaque (ODP) to establish contact and adhere to neighboring cells in a Ca²⁺-dependent manner. The cadherin cytoplasmic tails associate linker proteins, plakoglobin (PG), the plakophilins (PKP), and desmoplakin (DP). DP binds to keratin intermediate filaments (KIF) within the inner dense plaque (IDP), serving to tether the intermediate filaments to the plasma membrane. (Adapted with permission from Kottke et al. 2006.)In the mid 1970s, Skerrow and Matoltsy (Skerrow and Matoltsy 1974a; Skerrow and Matoltsy 1974b) advanced the field by isolating desmosomes using biochemical approaches (Bass-Zubek and Green 2007).These landmark studies provided a foundation for the Franke and Steinberg laboratories to characterize the transmembrane glycoproteins and cytoplasmic plaque proteins that linked the structure to the intermediate filament cytoskeleton, and to develop immunological tools for localizing specific components (Franke et al. 1981; Kapprell et al. 1985; Steinberg et al. 1987). Collectively, these and other studies shaped our current view of how desmosomal components are organized.The transmembrane glycoproteins, termed desmogleins and desmocollins (Garrod and Chidgey 2008), represent separate subfamilies of the cadherin superfamily of calcium dependent adhesion molecules. The extracellular domains of the desmogleins and desmocollins mediate adhesion, whereas the cytoplasmic tails of these cadherins associate with the desmosomal plaque proteins. The outer dense plaque consists of the cytoplasmic tails of the desmosomal cadherins, which bind to members of the armadillo and plakin family of linker proteins (Kowalczyk et al. 1994; Getsios et al. 2004b; Garrod and Chidgey 2008). Plakoglobin, a member of the armadillo family, binds directly to the cytoplasmic tails of both the desmogleins and the desmocollins (Wahl et al. 1996; Witcher et al. 1996). Desmoplakin, a member of the plakin family, interacts with both plakoglobin and another subgroup of armadillo family proteins, the plakophilins (Cowin and Burke 1996). Finally, the interaction between desmoplakin and the keratin filaments forms the inner dense plaque, tethering the cytoskeletal network to the adhesion complex (Fig. 1B) (Kowalczyk et al. 1994; Getsios et al. 2004b; Garrod and Chidgey 2008).The following sections of this article describe the structural and functional characteristics of the major desmosomal proteins. In addition, we discuss differences in tissue expression patterns of desmosomal proteins and the role of desmosomes in human disease. A comprehensive review of additional proteins found to regulate or associate with desmosomes is provided elsewhere (Holthofer et al. 2007) and discussion of desmosome dynamics is provided in Green et al. 2009.     

Self-avoidance and Tiling: Mechanisms of Dendrite and Axon Spacing

The spatial pattern of branches within axonal or dendritic arbors and the relative arrangement of neighboring arbors with respect to one another impact a neuron''s potential connectivity. Although arbors can adopt diverse branching patterns to suit their functions, evenly spread branches that avoid clumping or overlap are a common feature of many axonal and dendritic arbors. The degree of overlap between neighboring arbors innervating a surface is also characteristic within particular neuron types. The arbors of some populations of neurons innervate a target with a comprehensive and nonoverlapping “tiled” arrangement, whereas those of others show substantial territory overlap. This review focuses on cellular and molecular studies that have provided insight into the regulation of spatial arrangements of neurite branches within and between arbors. These studies have revealed principles that govern arbor arrangements in dendrites and axons in both vertebrates and invertebrates. Diverse molecular mechanisms controlling the spatial patterning of sister branches and neighboring arbors have begun to be elucidated.Axonal and dendritic arbors adopt complex and morphologically diverse shapes that influence neural connectivity and information processing. In this article we review anatomical and molecular studies that elucidate how the arrangements of branches within neuronal arbors are established during development (isoneuronal spacing) and how the relative spacing of arbors is determined when multiple neurons together innervate a defined territory (heteroneuronal spacing). Together these mechanisms ensure that arbors achieve functionally appropriate coverage of input or output territories.Isoneuronal and heteroneuronal processes display a variety of spacing arrangements, suggesting a diversity of underlying molecular mechanisms. Self-avoidance can occur between branches that arise from a single soma (Yau 1976; Kramer and Kuwada 1983; Kramer and Stent 1985), implying that neurons are able to discriminate “self,” which they avoid, from “nonself” arbors, with which they coexist (Kramer and Kuwada 1983). Similarly, arbors from different cells that share the same function and together innervate a defined territory can create a pattern of minimally overlapping neighboring dendritic or axonal fields, known as tiling. Such spacing mechanisms ensure that arbors maximize their spread across a territory while minimizing the redundancy with which the territory is innervated. In contrast, adhesive interactions between arbors can operate to maintain coherence of dendrites at specific targets (Zhu and Luo 2004), or to bundle functionally similar processes and possibly coordinate their activity (Campbell et al. 2009). Understanding how processes are patterned relative to one another can help to uncover the functional logic of neural circuit organization.Here we focus primarily on mechanisms of isoneuronal and heteroneuronal avoidance that result in complete and nonredundant innervation of sensory or synaptic space. Such mechanisms have been studied extensively in systems where neuronal arbors innervate a two-dimensional plane, such as the retina or body wall (Wassle et al. 1981; Perry and Linden 1982; Hitchcock 1989; Lin and Masland 2004; Fuerst et al. 2009; Kramer and Stent 1985; Grueber et al. 2003; Sugimura et al. 2003; Sagasti et al. 2005). However, the principles regulating process spacing in these regions likely also apply in three dimensions, most prominently where processes are segregated into nonoverlapping domains or columns (Huckfeldt et al. 2009). It is also notable that nonneuronal cell types might similarly engage in self-avoidance and form tiling arrangements, including leech comb cells (Jellies and Kristan 1991) and mammalian astrocytes (Bushong et al. 2002; Ogata and Kosaka 2002; Livet et al. 2007). Elucidating the mechanisms of process spacing during development is therefore relevant for understanding principles of tissue organization inside and outside of the nervous system.     

The Measure of Success: Constraints, Objectives, and Tradeoffs in Morphogen-mediated Patterning

A large, diverse, and growing number of strategies have been proposed to explain how morphogen gradients achieve robustness and precision. We argue that, to be useful, the evaluation of such strategies must take into account the constraints imposed by competing objectives and performance tradeoffs. This point is illustrated through a mathematical and computational analysis of the strategy of self-enhanced morphogen clearance. The results suggest that the usefulness of this strategy comes less from its ability to increase robustness to morphogen source fluctuations per se, than from its ability to overcome specific kinds of noise, and to increase the fraction of a morphogen gradient within which robust threshold positions may be established. This work also provides new insights into the longstanding question of why morphogen gradients show a maximum range in vivo.In recent years, much research on morphogen gradients has shifted from purely mechanistic questions—how gradients form and how morphogens signal—to strategic ones—how gradients perform well in the face of various kinds of constraints and perturbations. Forty years ago, Francis Crick was among the first to call attention to constraints that morphogens face, noting that the time required to spread a signal by random transport through a tissue varies with the square of distance (Crick 1970). Using order-of-magnitude calculations, he argued that observed biological maxima for morphogen-mediated patterning were just about where they should be if morphogen signals spread by aqueous diffusion.Although the idea that diffusion time is what limits the sizes of morphogen gradients remains untested, Crick''s work established a precedent of seeking explanations for developmental processes in terms of constraints imposed by the physical world. In the area of biological pattern formation, continued interest in how real-world limits constrain mechanisms has led many current investigators to focus on matters of robustness, the engineering term that describes the relative insensitivity of a system''s behavior to perturbations it may be expected to encounter. With respect to morphogen gradients, most work has focused on parametric robustness, i.e., insensitivity to parameter values (e.g., the dosage of genes, levels, or rate constants of enzymes [Eldar et al. 2002; Eldar et al. 2003; Eldar et al. 2004; Bollenbach et al. 2005; Shimmi et al. 2005; White et al. 2007]). Some investigators have also focused on the “precision” of morphogen gradients, which may be understood as robustness to the causes and effects of natural variation among individuals in a population (Houchmandzadeh et al. 2002; Gregor et al. 2007; Tostevin et al. 2007; Bollenbach et al. 2008; Emberly 2008).Remarkably, after hardly a decade of intensive study of such questions, we find ourselves awash in a sea of diverse and intriguing mechanisms for conferring one or another type of robustness on morphogen-mediated patterning. Mechanisms that operate at the level of gradient formation include self-enhanced morphogen degradation (Eldar et al. 2003), facilitated transport (Eldar et al. 2002; Shimmi et al. 2005), serial transcytosis (Bollenbach et al. 2005), presteady state patterning (Bergmann et al. 2007), and competition between morphogens for binding to inhibitors (Ben-Zvi et al. 2008). Mechanisms that operate at the level of morphogen detection and interpretation include morphogenetic apoptosis (Adachi-Yamada and O''Connor 2002), cell rearrangement (Ashe and Briscoe 2006), integration of signals from multiple morphogens (McHale et al. 2006; Morishita and Iwasa 2008), and various types of local cell-to-cell signaling (e.g., Amonlirdviman et al. 2005).Why so many strategies? Biologists are often quick to ascribe multiplicity to redundancy, but the perspective of engineering suggests a different view. Most engineers accept the “no free lunch” principle (also referred to as “conservation of fragility”), which states that any mechanism that increases robustness in one setting (i.e., to one type of perturbation, or with respect to one type of output) always compromises it in another. The fact that every strategy comes at a price has been offered as an explanation for the seemingly inescapable fragility of highly engineered, modern technology (Carlson and Doyle 2002). By building complex machines that resist everything we think of, we inevitably create susceptibilities to the things we neglected. Although biology is not the result of human engineering, we have no reason to believe that natural selection can circumvent the limits that engineers confront.In a world of no free lunch, one must evaluate a strategy not just by what it is good for, but the “price” of using it. With regard to morphogen-mediated patterning, it is reasonable to suggest that diverse strategies exist because each comes at a different price. If so, achieving meaningful biological understanding requires that we engage in a sort of cost-benefit analysis, in which each strategy is evaluated in the context of the performance objectives of the organism and constraints of the physical world. This is a tall order, as there is a great deal we still do not know about the performance needs of developing organisms (for example, for all the work performed so far on morphogen gradient robustness, we still know little about the magnitudes of the perturbations that need to be withstood). Nevertheless, there is no reason not to get started, as even through the early investigation of hard questions, one commonly learns useful things.     

Dynamic Transbilayer Lipid Asymmetry

Cells have thousands of different lipids. In the plasma membrane, and in membranes of the late secretory and endocytotic pathways, these lipids are not evenly distributed over the two leaflets of the lipid bilayer. The basis for this transmembrane lipid asymmetry lies in the fact that glycerolipids are primarily synthesized on the cytosolic and sphingolipids on the noncytosolic surface of cellular membranes, that cholesterol has a higher affinity for sphingolipids than for glycerolipids. In addition, P4-ATPases, “flippases,” actively translocate the aminophospholipids phosphatidylserine and phosphatidylethanolamine to the cytosolic surface. ABC transporters translocate lipids in the opposite direction but they generally act as exporters rather than “floppases.” The steady state asymmetry of the lipids can be disrupted within seconds by the activation of phospholipases and scramblases. The asymmetric lipid distribution has multiple implications for physiological events at the membrane surface. Moreover, the active translocation also contributes to the generation of curvature in the budding of transport vesicles.A lipid bilayer consisting of phosphatidylcholine (PC) with one saturated and one unsaturated acyl chain is stable, flexible, and semipermeable. It is the simplest model of a biomembrane. In such membranes, PC with a spin label on its choline headgroup diffused rapidly in the plane of the membrane with a diffusion coefficient of 1.8 µm²/sec (Devaux and McConnell 1972). In contrast, PC movement between leaflets, “flip-flop,” was slow with a half-time of >6 h at 30°C (Kornberg and McConnell 1971). Similar half-times for PC flip-flop were measured in erythrocyte membranes, a mammalian plasma membrane with a complex lipid composition (Rousselet et al. 1976; Renooij and Van Golde 1977; van Meer et al. 1980). Interestingly, the erythrocyte membrane maintains an asymmetric lipid distribution across the lipid bilayer with all of its phosphatidylserine (PS) and most of its phosphatidylethanolamine (PE) in the cytosolic leaflet (Bretscher 1972; Verkleij et al. 1973). A critical discussion of these early data and the techniques used can be found in (Op den Kamp 1979).It was then observed that the enrichment of aminophospholipids in the cytosolic leaflet is maintained by an ATP-consuming translocator that flips these lipids from the outer leaflet across the lipid bilayer (Seigneuret and Devaux 1984). The flippase was later identified as a P4-ATPase (Tang et al. 1996; Soupene and Kuypers 2006). Around the same time it was found that an ABC transporter, ABCB4, was involved in transporting PC into the bile (Smit et al. 1993), and studies on the closely related ABCB1 proved that these transporters can translocate lipids across the plasma membrane onto acceptors in the extracellular space (van Helvoort et al. 1996). Finally, evidence was provided for passive, bidirectional movement of lipids across the ER membrane and under some conditions across the plasma membrane, in which cases the responsible proteins have not yet been unequivocally identified (Sanyal and Menon 2009; Bevers and Williamson 2010). Thus, we now have a general picture of how lipid asymmetry is generated, maintained, and disrupted. However, there are still important gaps in our knowledge. For example, the transbilayer orientation of the sterols that make up one-third of the lipids in eukaryotic plasma membranes has still not been resolved satisfactorily. Moreover, we do not understand mechanistically how translocators and exporters work and how their activity is regulated.     

Origin and Evolution of the Self-Organizing Cytoskeleton in the Network of Eukaryotic Organelles

The eukaryotic cytoskeleton evolved from prokaryotic cytomotive filaments. Prokaryotic filament systems show bewildering structural and dynamic complexity and, in many aspects, prefigure the self-organizing properties of the eukaryotic cytoskeleton. Here, the dynamic properties of the prokaryotic and eukaryotic cytoskeleton are compared, and how these relate to function and evolution of organellar networks is discussed. The evolution of new aspects of filament dynamics in eukaryotes, including severing and branching, and the advent of molecular motors converted the eukaryotic cytoskeleton into a self-organizing “active gel,” the dynamics of which can only be described with computational models. Advances in modeling and comparative genomics hold promise of a better understanding of the evolution of the self-organizing cytoskeleton in early eukaryotes, and its role in the evolution of novel eukaryotic functions, such as amoeboid motility, mitosis, and ciliary swimming.The eukaryotic cytoskeleton organizes space on the cellular scale and this organization influences almost every process in the cell. Organization depends on the mechanochemical properties of the cytoskeleton that dynamically maintain cell shape, position organelles, and macromolecules by trafficking, and drive locomotion via actin-rich cellular protrusions, ciliary beating, or ciliary gliding. The eukaryotic cytoskeleton is best described as an “active gel,” a cross-linked network of polymers (gel) in which many of the links are active motors that can move the polymers relative to each other (Karsenti et al. 2006). Because prokaryotes have only cytoskeletal polymers but lack motor proteins, this “active gel” property clearly sets the eukaryotic cytoskeleton apart from prokaryotic filament systems.Prokaryotes contain elaborate systems of several cytomotive filaments (Löwe and Amos 2009) that share many structural and dynamic features with eukaryotic actin filaments and microtubules (Löwe and Amos 1998; van den Ent et al. 2001). Prokaryotic cytoskeletal filaments may trace back to the first cells and may have originated as higher-order assemblies of enzymes (Noree et al. 2010; Barry and Gitai 2011). These cytomotive filaments are required for the segregation of low copy number plasmids, cell rigidity and cell-wall synthesis, cell division, and occasionally the organization of membranous organelles (Komeili et al. 2006; Thanbichler and Shapiro 2008; Löwe and Amos 2009). These functions are performed by dynamic filament-forming systems that harness the energy from nucleotide hydrolysis to generate forces either via bending or polymerization (Löwe and Amos 2009; Pilhofer and Jensen 2013). Although the identification of actin and tubulin homologs in prokaryotes is a major breakthrough, we are far from understanding the origin of the structural and dynamic complexity of the eukaryotic cytoskeleton.Advances in genome sequencing and comparative genomics now allow a detailed reconstruction of the cytoskeletal components present in the last common ancestor of eukaryotes. These studies all point to an ancestrally complex cytoskeleton, with several families of motors (Wickstead and Gull 2007; Wickstead et al. 2010) and filament-associated proteins and other regulators in place (Jékely 2003; Richards and Cavalier-Smith 2005; Rivero and Cvrcková 2007; Chalkia et al. 2008; Eme et al. 2009; Fritz-Laylin et al. 2010; Eckert et al. 2011; Hammesfahr and Kollmar 2012). Genomic reconstructions and comparative cell biology of single-celled eukaryotes (Raikov 1994; Cavalier-Smith 2013) allow us to infer the cellular features of the ancestral eukaryote. These analyses indicate that amoeboid motility (Fritz-Laylin et al. 2010; although, see Cavalier-Smith 2013), cilia (Cavalier-Smith 2002; Mitchell 2004; Jékely and Arendt 2006; Satir et al. 2008), centrioles (Carvalho-Santos et al. 2010), phagocytosis (Cavalier-Smith 2002; Jékely 2007; Yutin et al. 2009), a midbody during cell division (Eme et al. 2009), mitosis (Raikov 1994), and meiosis (Ramesh et al. 2005) were all ancestral eukaryotic cellular features. The availability of functional information from organisms other than animals and yeasts (e.g., Chlamydomonas, Tetrahymena, Trypanosoma) also allow more reliable inferences about the ancestral functions of cytoskeletal components (i.e., not only their ancestral presence or absence) and their regulation (Demonchy et al. 2009; Lechtreck et al. 2009; Suryavanshi et al. 2010).The ancestral complexity of the cytoskeleton in eukaryotes leaves a huge gap between prokaryotes and the earliest eukaryote we can reconstruct (provided that our rooting of the tree is correct) (Cavalier-Smith 2013). Nevertheless, we can attempt to infer the series of events that happened along the stem lineage, leading to the last common ancestor of eukaryotes. Meaningful answers will require the use of a combination of gene family history reconstructions (Wickstead and Gull 2007; Wickstead et al. 2010), transition analyses (Cavalier-Smith 2002), and computer simulations relevant to cell evolution (Jékely 2008).     

SLAM-Family Receptors: Immune Regulators with or without SAP-Family Adaptors

The signaling lymphocytic activation molecule (SLAM) family of receptors and the SLAM-associated protein (SAP) family of intracellular adaptors are expressed in immune cells. By way of their cytoplasmic domain, SLAM-related receptors physically associate with SAP-related adaptors. Evidence is accumulating that the SLAM and SAP families play crucial roles in multiple immune cell types. Moreover, the prototype of the SAP family, that is SAP, is mutated in a human immunodeficiency, X-linked lymphoproliferative (XLP) disease. In the presence of SAP-family adaptors, the SLAM family usually mediates stimulatory signals that promote immune cell activation or differentiation. In the absence of SAP-family adaptors, though, the SLAM family undergoes a “switch-of-function,” thereby mediating inhibitory signals that suppress immune cell functions. The molecular basis and significance of this mechanism are discussed herein.Immune cells undergo differentiation and, once mature, are activated through the integrated actions of many molecules, including cell surface receptors and intracellular signaling effectors. Whereas some of these molecules have “primary” roles in the immune response, others have secondary, albeit still critical, functions in this process. For example, differentiation and activation of B cells are strictly dependent on the function of the B-cell receptor (BCR) and its intracellular effectors. Other receptors present on B cells, such as CD19 and CD40, influence B-cell functions in critical ways, by modulating BCR-triggered signals (Cambier et al. 1994).There is accumulating evidence that the signaling lymphocytic activation molecule (SLAM) family of receptors plays important roles in immunity (Schwartzberg et al. 2009; Ma et al. 2007; Veillette et al. 2007; Veillette 2006b; Veillette 2006a). This class of receptors provides key effects in multiple immune cell types. Recent data indicate that SLAM-family receptors can either promote or inhibit the functions of primary activating receptors (Cruz-Munoz et al. 2009; Dong et al. 2009). These alternative activities are controlled by whether or not SLAM-related receptors are coexpressed with members of the SLAM-associated protein (SAP) family of intracellular adaptor molecules. The functions and mechanisms of action of the SLAM and SAP families are reviewed herein.     

Effects of Membrane Trafficking on Signaling by Receptor Tyrosine Kinases

The intracellular trafficking machinery contributes to the spatial and temporal control of signaling by receptor tyrosine kinases (RTKs). The primary role in this process is played by endocytic trafficking, which regulates the localization of RTKs and their downstream effectors, as well as the duration and the extent of their activity. The key regulatory points along the endocytic pathway are internalization of RTKs from the plasma membrane, their sorting to degradation or recycling, and their residence in various endosomal compartments. Here I will review factors and mechanisms that modulate RTK signaling by (1) affecting receptor internalization, (2) regulating the balance between degradation and recycling of RTK, and (3) compartmentalization of signals in endosomes and other organelles. Cumulatively, these mechanisms illustrate a multilayered control of RTK signaling exerted by the trafficking machinery.At the cellular level, receptor tyrosine kinases (RTKs) need to be properly localized to function as signal-receiving and signal-transmitting devices (Lemmon and Schlessinger 2010). To receive signals (i.e., to bind extracellular ligands), RTKs have to be exposed at the surface of the plasma membrane. To transmit signals after ligand binding by RTKs, appropriate signaling components have to be available within intracellular compartments: in the cytoplasm, in association with membrane-bound organelles and in the cell nucleus. Importantly, the intracellular distribution of RTKs and their associated partners is not static but undergoes dynamic changes in different phases of signaling, as reflected for example by endocytic internalization of activated RTKs (Scita and Di Fiore 2010). Therefore, to function properly, the whole RTK signaling machinery within the cell has to be organized and tightly controlled both in space and in time. This organization and control are ensured by intracellular trafficking machineries, mainly by membrane transport systems such as endocytosis and secretion but also by other distribution systems (e.g., responsible for nucleocytoplasmic shuttling of proteins).Recent years have brought increasing evidence that intracellular membrane trafficking, in particular endocytic internalization, degradation, and recycling, can profoundly affect the signaling properties of RTKs (Mukherjee et al. 2006; Abella and Park 2009; Lemmon and Schlessinger 2010; Scita and Di Fiore 2010; Grecco et al. 2011; Sigismund et al. 2012). The changes in the amounts of RTKs at the cell surface can alter the cellular responses when ligands are abundant (Grecco et al. 2011). In turn, the presence of a given RTK at the plasma membrane is determined by the rates of three trafficking processes: delivery of newly synthesized molecules by the secretory pathway, their internalization (occurring for both ligand-bound and ligand-free molecules), and endocytic recycling. Although the molecular details concerning the regulation of RTK delivery to the plasma membrane are not well known, numerous studies document various mechanisms by which internalization and recycling of RTKs can be modulated, thus affecting the signaling outputs (Le Roy and Wrana 2005). In addition to the regulation of RTKs at the cell surface, trafficking processes control the intracellular fate of endocytosed RTKs. Following internalization, RTKs can be either targeted for lysosomal degradation, or recycled back to the plasma membrane (Mukherjee et al. 2006; Abella and Park 2009; Scita and Di Fiore 2010). The first route results in the termination of signaling, whereas the second allows for sustained signaling if the ligand is available. Usually degradation and recycling of a given RTK can occur simultaneously but the balance between them is crucial to determine the net signaling output. Again, the molecular mechanisms that can shift the fate of internalized RTKs between degradation and recycling, thus changing RTK signaling, have begun to emerge in recent years (Polo and Di Fiore 2006; von Zastrow and Sorkin 2007; Sorkin and von Zastrow 2009; Sigismund et al. 2012). Finally, in contrast to an early view that only RTKs present at the plasma membrane are signaling competent, it is now accepted that in many cases activated RTKs can emit signals also after internalization into intracellular compartments (Miaczynska et al. 2004b; Miaczynska and Bar-Sagi 2010; Platta and Stenmark 2011). In some cell types (e.g., in neurons), such “signaling endosomes” are crucial for signal propagation within the cell and for the final cellular response. Moreover, endosomes can serve as platforms for amplification and compartmentalization of signals emitted by RTKs (Sadowski et al. 2009; Platta and Stenmark 2011).In this article, I will review factors and mechanisms that modulate RTK signaling by (1) affecting receptor internalization, (2) regulating the balance between degradation and recycling of RTK, and (3) compartmentalization of signals in endosomes and other organelles. As the membrane trafficking system of a cell is highly interconnected and can be considered a global dynamic continuum, it is important to note that often one primary alteration at a given stage of RTK trafficking may affect other transport steps or compartments, thus causing generalized changes in the intracellular routing and signaling of RTKs.     

Integrin Structure,Activation, and Interactions

Integrins are large, membrane-spanning, heterodimeric proteins that are essential for a metazoan existence. All members of the integrin family adopt a shape that resembles a large “head” on two “legs,” with the head containing the sites for ligand binding and subunit association. Most of the receptor dimer is extracellular, but both subunits traverse the plasma membrane and terminate in short cytoplasmic domains. These domains initiate the assembly of large signaling complexes and thereby bridge the extracellular matrix to the intracellular cytoskeleton. To allow cells to sample and respond to a dynamic pericellular environment, integrins have evolved a highly responsive receptor activation mechanism that is regulated primarily by changes in tertiary and quaternary structure. This review summarizes recent progress in the structural and molecular functional studies of this important class of adhesion receptor.The name “integrin” was suggested for an integral membrane protein complex first characterized in 1986 (Tamkun et al. 1986). The name was devised because the protein identified linked the extracellular matrix to the cytoskeleton (early developments in this field have been well described [Hynes 2004]). In the 25 years since that first characterization, a vast amount of work has been performed, with consequent increased understanding. The essential role of integrins in tissue organization and cell development, their signal transduction mechanisms (from outside to in and inside to out!), and their potential as therapeutic targets is now established. In this article, we provide an overview of the structure of integrins, the conformational changes that determine activation state, and the mechanisms of ligand binding.     

Morphogen Gradient Formation

How morphogen gradients are formed in target tissues is a key question for understanding the mechanisms of morphological patterning. Here, we review different mechanisms of morphogen gradient formation from theoretical and experimental points of view. First, a simple, comprehensive overview of the underlying biophysical principles of several mechanisms of gradient formation is provided. We then discuss the advantages and limitations of different experimental approaches to gradient formation analysis.How a multicellular organism develops from a single fertilized cell has fascinated people throughout history. By looking at chick embryos of different developmental stages, Aristotle first noted that development is characterized by growing complexity and organization of the embryo (Balme 2002). During the 19th century, two events were recognized as key in development: cell proliferation and differentiation. Driesch first noted that to form organisms with correct morphological pattern and size, these processes must be controlled at the level of the whole organism. When he separated two sea urchin blastomeres, they produced two half-sized blastula, showing that cells are potentially independent, but function together to form a whole organism (Driesch 1891, 1908). Morgan noted the polarity of organisms and that regeneration in worms occurs with different rates at different positions. This led him to postulate that regeneration phenomena are influenced by gradients of “formative substances” (Morgan 1901).The idea that organisms are patterned by gradients of form-providing substances was explored by Boveri and Hörstadius to explain the patterning of the sea urchin embryo (Boveri 1901; Hörstadius 1935). The discovery of the Spemann organizer, i.e., a group of dorsal cells that when grafted onto the opposite ventral pole of a host gastrula induce a secondary body axis (Spemann and Mangold 1924), suggested that morphogenesis results from the action of signals that are released from localized groups of cells (“organizing centers”) to induce the differentiation of the cells around them (De Robertis 2006). Child proposed that these patterning “signals” represent metabolic gradients (Child 1941), but the mechanisms of their formation, regulation, and translation into pattern remained elusive.In 1952, Turing showed that chemical substances, which he called morphogens (to convey the idea of “form producers”), could self-organize into spatial patterns, starting from homogenous distributions (Turing 1952). Turing’s reaction–diffusion model shows that two or more morphogens with slightly different diffusion properties that react by auto- and cross-catalyzing or inhibiting their production, can generate spatial patterns of morphogen concentration. The reaction–diffusion formalism was used to model regeneration in hydra (Turing 1952), pigmentation of fish (Kondo and Asai 1995; Kondo 2002), and snails (Meinhardt 2003).At the same time that Turing showed that pattern can self-organize from the production, diffusion, and reaction of morphogens in all cells, the idea that morphogens are released from localized sources (“organizers” à la Spemann) and form concentration gradients was still explored. This idea was formalized by Wolpert with the French flag model for generation of positional information (Wolpert 1969). According to this model, morphogen is secreted from a group of source cells and forms a gradient of concentration in the target tissue. Different target genes are expressed above distinct concentration thresholds, i.e., at different distances to the source, hence generating a spatial pattern of gene expression (Fig. 1C).Open in a separate windowFigure 1.Tissue geometry and simplifications. (A) Gradients in epithelia (left) and mesenchymal tissues (right). Because of symmetry considerations, one row of cells (red outline) is representative for the whole gradient. (B) Magnified view of the red row of cells shown in A. Cells with differently colored nuclei (brown, orange, and blue) express different target genes. (C) A continuum model in which individual cells are ignored and the concentration is a function of the positions x. The morphogen activates different target genes above different concentration thresholds (brown and orange).Experiments in the 1970s and later confirmed that tissues are patterned by morphogen gradients. Sander showed that a morphogen released from the posterior cytoplasm specifies anterioposterior position in the insect egg (Sander 1976). Chick wing bud development was explained by a morphogen gradient emanating from the zone of polarizing activity to specify digit positions (Saunders 1972; Tickle, et al. 1975; Tickle 1999). The most definitive example of a morphogen was provided with the identification of Bicoid function in the Drosophila embryo (Nüsslein-Volhard and Wieschaus 1980; Frohnhöfer and Nüsslein-Volhard 1986; Nüsslein-Volhard et al. 1987) and the visualization of its gradient by antibody staining (Driever and Nüsslein-Volhard 1988b, 1988a; reviewed in Ephrussi and St Johnston 2004). Since then, many examples of morphogen gradients acting in different organs and species have been found.In an attempt to understand pattern formation in more depth, quantitative models of gradient formation have been developed. An early model by Crick shows that freely diffusing morphogen produced in a source cell and destroyed in a “sink” cell at a distance would produce a linear gradient in developmentally relevant timescales (Crick 1970). Today, it is known that a localized “sink” is not necessary for gradient formation: Gradients can form if all cells act as sinks and degrade morphogen, or even if morphogen is not degraded at all. Here, we review different mechanisms of gradient formation, the properties of these gradients, and the implications for patterning. We discuss the theory behind these mechanisms and the supporting experimental data.     

Architecture of the Mammalian Golgi

Since its first visualization in 1898, the Golgi has been a topic of intense morphological research. A typical mammalian Golgi consists of a pile of stapled cisternae, the Golgi stack, which is a key station for modification of newly synthesized proteins and lipids. Distinct stacks are interconnected by tubules to form the Golgi ribbon. At the entrance site of the Golgi, the cis-Golgi, vesicular tubular clusters (VTCs) form the intermediate between the endoplasmic reticulum and the Golgi stack. At the exit site of the Golgi, the trans-Golgi, the trans-Golgi network (TGN) is the major site of sorting proteins to distinct cellular locations. Golgi functioning can only be understood in light of its complex architecture, as was revealed by a range of distinct electron microscopy (EM) approaches. In this article, a general concept of mammalian Golgi architecture, including VTCs and the TGN, is described.In 1898 Camillo Golgi was the first to visualize, describe, and ultimately name the Golgi complex. Using a histochemical impregnation method causing the reduction and deposition of silver, he defined the Golgi in neuronal cells as a reticular apparatus stained by the “black reaction” (Golgi 1898). In the 1950s, the first ultrastructural images of the Golgi were revealed using the then newly developed electron microscope (EM) (Dalton 1954; Farquhar and Rinehart 1954; Sjostrand and Hanzon 1954; Dalton and Felix 1956), reviewed by Farquhar and Palade (1981). In 1961, the thiamine pyrophosphatase reaction developed by Novikoff and Goldfischer allowed cytochemical labeling of Golgi membranes, which revealed the ubiquitous cellular distribution of this organelle (Novikoff and Goldfischer 1961). In the many years of ultrastructural research that have followed, the visualization of the Golgi has gone hand-in-hand with the developing EM techniques.The intriguing structural complexity of the Golgi has made it one of the most photographed organelles in the cell. However, a full understanding of Golgi architecture is hard to deduce from the ultrathin (70–100 nm) sections used in standard transmission EM preparations. Rambourg and Clermont (1974) were the first to investigate the Golgi in three dimensions (3D), using stereoscopy (Rambourg 1974). In this approach a “thick” (150–200 nm), EM section is photographed at two distinct angles, after which the pairs of photographs are viewed with a stereoscope. Over the years, stereoscopy was applied to a variety of cells and has greatly contributed to our current understanding of Golgi architecture (Lindsey and Ellisman 1985; Rambourg and Clermont 1990; Clermont et al. 1994; Clermont et al. 1995). An alternative approach to study 3D structure is serial sectioning, by which a series of adjacent (serial) thin sections are collected. The Golgi can be followed throughout these sections and be constructed into a 3D model (Beams and Kessel 1968; Dylewski et al. 1984; Rambourg and Clermont 1990). In the nineties, 3D-EM was boosted by the introduction of high-voltage, dual axis 3D electron tomography (Ladinsky et al. 1999; Koster and Klumperman 2003; Marsh 2005; Marsh 2007; Noske et al. 2008), which allows the analysis of sections of up to 3–4 µm with a 4–6 nm resolution in the z-axis. The sections are photographed in a tilt series of different angles, which are reconstructed into a 3D tomogram that allows one to “look beyond” a given structure and reveals how it relates to other cellular compartments.Membranes with a similar appearance can differ in protein content and function. These differences are revealed by protein localization techniques. Therefore, in addition to the “classical” EM techniques providing ultrastructural details, EM methods that determine protein localization within the context of the cellular morphology have been crucial to further our understanding on the functional organization of the Golgi. For example, by enzyme-activity-based cytochemical staining the cis-to-trans-polarity in the distribution of Golgi glycosylation enzymes was discovered, reviewed by Farquhar and Palade (1981), which was key to understanding the functional organization of the Golgi stack in protein and lipid glycosylation. With the development of immunoEM methods, using antibodies, the need for enzyme activity for protein localization was overcome. This paved the way for the localization of a wide variety of proteins, such as the cytoplasmic coat complexes associated with the Golgi (Rabouille and Klumperman 2005).A logical next step in EM-based imaging of the Golgi would be to combine protein localization with 3D imaging, but this is technically challenging. A number of protocols enabling protein localization in 3D have recently been described (Trucco et al. 2004; Grabenbauer et al. 2005; Gaietta et al. 2006; Zeuschner et al. 2006; Meiblitzer-Ruppitsch et al. 2008), but these have only been applied in a limited manner to Golgi studies. Another approach that holds great potential for Golgi research is correlative microscopy (CLEM). Live cell imaging of fluorescent proteins has revolutionized cell biology by the real time visualization of dynamic events. However, live cell imaging does not reveal membrane complexity. By CLEM, live cells are first viewed by light microscopy and then prepared for EM (Mironov et al. 2008; van Rijnsoever et al. 2008). When coupled with the recent introduction of super resolution light microscopy techniques for real time imaging, the combination with EM for direct correlation with ultrastructural resolution has great potential (Hell 2009; Lippincott-Schwartz and Manley 2009).The 100th anniversary of the discovery of the Golgi, in 1998, triggered a wave of reviews on this organelle, including those focusing on Golgi architecture (Rambourg 1997; Farquhar and Palade 1998). More recent reviews that describe Golgi structure in great detail are provided by Marsh (2005) and Hua (2009). In this article, the most recent insights in mammalian Golgi architecture as revealed by distinct EM approaches are integrated into a general concept.     

Molecular Basis of the Core Structure of Tight Junctions

The morphological feature of tight junctions (TJs) fits well with their functions. The core of TJs is a fibril-like proteinaceous structure within the lipid bilayer, the so-called TJ strands. TJ strands in apposing plasma membranes associate with each other to eliminate the intercellular space. A network of paired TJ strands generates a continuous belt that circumscribes each cell to establish the diffusion barrier to the solutes in the paracellular pathway throughout the cellular sheet. Identification and characterization of TJ-associated proteins during the last two decades has unveiled the nature of TJ strands and how they are spatially organized. The interplay between integral membrane proteins, claudins, and cytoplasmic plaque proteins, ZO-1/ZO-2, is critical for TJ formation and function.Tight junctions (TJs) are fascinating structures in terms of their function and morphology. In 1963, using ultrathin-section electron microscopy, Farquhar and Palade described the fine structure of TJs together with adherens junctions (AJs) and desmosomes at the most luminal side of the lateral membrane (Farquhar and Palade 1963). In addition, they demonstrated insightfully that TJs function as permeability seals for mass tracers. Indeed, the structure of TJs observed in electron microscopy indicates that TJs could physically restrict the leak of solutes through the intercellular space. However, physiological studies at the same time revealed that solute transport occurred via the intercellular space in a variety of epithelial cells. A resolution of these different views of TJ function comprises the current concept that the TJ regulates the diffusion of solutes with size and charge selectivity and that it is functionally different in physiologically diverse epithelial cell types (Powell 1981; Anderson and Cereijido 2001). To understand the molecular mechanism controlling TJ structure and function, it is important to determine their molecular composition and organization.Although purification of TJs is difficult, Stevenson and Goodenough developed an isolation method for a TJ-enriched plasma membrane fraction from rodent liver. They discovered the first TJ-associated protein, ZO-1, in 1986 by generating monoclonal antibodies against this fraction (Stevenson et al. 1986). Since then, many molecular components of TJs have been identified using immunological approaches or searches for binding proteins with known molecules, which have enabled detailed molecular cell biological analyses of TJs. Among the TJ-associated proteins, the claudin family of membrane proteins identified in 1998 by the Tsukita group are key molecules in the architecture and barrier function of TJs (Furuse et al. 1998a). Functional analyses of claudins have allowed remarkable progress in the development of a comprehensive understanding of the molecular basis of the ultrastructure and physiological characteristics of TJs (Van Itallie and Anderson 2006; Furuse and Tsukita 2006; Angelow et al. 2008). In addition, the cytoplasmic plaque proteins associated with TJs are important in regulating TJ architecture (Guillemot et al. 2008).In this article, we present the molecular basis for the core structure of TJs based on recent progress in functional analyses of TJ-associated proteins. The current molecular basis of TJ physiology is covered in detail in Anderson and Van Itallie (2009).     

Membrane Organization and Dynamics in Cell Polarity

The establishment and maintenance of cell polarity is important to a wide range of biological processes ranging from chemotaxis to embryogenesis. An essential feature of cell polarity is the asymmetric organization of proteins and lipids in the plasma membrane. In this article, we discuss how polarity regulators such as small GTP-binding proteins and phospholipids spatially and kinetically control vesicular trafficking and membrane organization. Conversely, we discuss how membrane trafficking contributes to cell polarization through delivery of polarity determinants and regulators to the plasma membrane.Cell polarity is essential in most if not all eukaryotes for their development and physiological functions at the tissue and organism level. Although there are significant differences in gross morphology and function among various tissues and organisms, at the cellular level, the establishment and maintenance of cell polarity tend to follow common themes.A basic feature of cell polarity is the asymmetric organization of the plasma membrane (see McCaffrey and Macara 2009; Nelson 2009). This is mostly achieved through membrane trafficking along cytoskeleton tracks under the control of signaling molecules. In general, membrane trafficking occurs through sequential budding, transport, and fusion of vesicles from donor membranes to acceptor membranes (for recent reviews, see Bonifacino and Glick 2004; Cai et al. 2007). During budding, protein complexes interact with phospholipids to induce membrane curvature and generate vesicular carriers that capture different cargos from the donor compartments. After vesicles form, they are delivered to their acceptor compartments, most often along the cytoskeletons. Vesicle fusion at the acceptor membrane is mediated by the assembly of SNARE (soluble N-ethylmaleimide-sensitive fusion protein attachment protein receptors) complexes. Before membrane fusion, proteins or protein complexes tether the vesicles to the acceptor membranes and likely promote SNARE assembly. The Arf and Rab family of small GTPases are localized to different membrane compartments and regulate various stages of membrane trafficking.Polarized distribution of proteins at the plasma membrane often results from a balance of vesicle delivery and fusion with the plasma membrane (“exocytosis”), two-dimensional spread through the plasma membrane (“diffusion”), and internalization and membrane recycling (“endocytosis”). There are two main layers of regulation that control polarized protein transport and incorporation to the plasma membrane. The first involves sorting at the trans-Golgi network (TGN) and endosomal compartments, such as the recycling endosomes. Protein sorting involves recognition of sorting signals in the cargo proteins by the adaptor protein (AP) complexes. There are a number of different AP complexes, and each is localized to different membrane compartments and captures distinct sets of cargo proteins before targeting to their correct destination. Protein sorting before delivery to different domains of the plasma membrane has been best characterized in epithelial cells, which have distinctive basolateral and apical domains separated by junctional complexes. This layer of regulation has been discussed in a recent review (Mellman and Nelson 2008) and is further discussed by Nelson (Nelson 2009), so it will not be discussed further here. The second layer of regulation of membrane protein polarization is through the polarized tethering and docking of vesicles at specific domains of the plasma membrane (Fig. 1). Tethering proteins (i.e., the exocyst) target secretory vesicles to specific domains of the plasma membrane and SNARE assembly eventually drives membrane fusion. Proteins at the plasma membrane can be retrieved back into the cell via endocytosis. These proteins are internalized via clathrin-coated pits, and transported through different endosomal compartments either for degradation in the lysosomes or for recycling back to the plasma membrane. The endosomal compartment that mediates the transport of internalized plasma membrane proteins back to the cell surface is called the “recycling endosome.” Recycling endosomes are major sources of cargo destined to the plasma membrane for exocytosis in many types of cells.Open in a separate windowFigure 1.Membrane trafficking to the plasma membrane. Schematic of the endocytic and exocytic routes involving trans-Golgi network (TGN), endosomal compartments, and the plasma membrane. During exocytosis, cargo leaves the TGN or recycling endosomes in vesicular carriers to the plasma membrane. Once on the membrane, proteins can be internalized and transported to early endosomes, and then either travel through late endosomes to the lysosome to be degraded or return to the plasma membrane through the recycling endosomes. Early endosomes may serve as sorting stations for the next stages of cargo transport.Signaling molecules such as the Rho family of small GTPases spatially and kinetically regulate membrane trafficking during cell polarization (see McCaffrey and Macara 2009; Slaughter et al. 2009). Reversely, vesicular trafficking is required for the polarized deposition and accrual of these regulators. In the first part of this article, we examine the membrane organization and dynamics of cell polarity, focusing on the polarized tethering and docking of vesicles at the plasma membrane. We highlight key components and regulators of polarized exocytosis including the exocyst, small GTPases, and phospholipids. We also use different organisms and systems to show analogous mechanisms during cell polarization. In the second part of this article, we focus on the aforementioned reciprocal effects of cell polarity and membrane trafficking using two representative examples, one from yeast (Cdc42 polarization) and one in mammalian epithelial cells (E-cadherin trafficking).     

Gap Junctions

Gap junctions are aggregates of intercellular channels that permit direct cell–cell transfer of ions and small molecules. Initially described as low-resistance ion pathways joining excitable cells (nerve and muscle), gap junctions are found joining virtually all cells in solid tissues. Their long evolutionary history has permitted adaptation of gap-junctional intercellular communication to a variety of functions, with multiple regulatory mechanisms. Gap-junctional channels are composed of hexamers of medium-sized families of integral proteins: connexins in chordates and innexins in precordates. The functions of gap junctions have been explored by studying mutations in flies, worms, and humans, and targeted gene disruption in mice. These studies have revealed a wide diversity of function in tissue and organ biology.Gap junctions are clusters of intercellular channels that allow direct diffusion of ions and small molecules between adjacent cells. The intercellular channels are formed by head-to-head docking of hexameric assemblies (connexons) of tetraspan integral membrane proteins, the connexins (Cx) (Goodenough et al. 1996). These channels cluster into polymorphic maculae or plaques containing a few to thousands of units (Fig. 1). The close membrane apposition required to allow the docking between connexons sterically excludes most other membrane proteins, leaving a narrow ∼2 nm extracellular “gap” for which the junction is named (Fig. 2). Gap junctions in prechordates are composed of innexins (Phelan et al. 1998; Phelan 2005). In chordates, connexins arose by convergent evolution (Alexopoulos et al. 2004), to expand by gene duplication (Cruciani and Mikalsen 2007) into a 21-member gene family. Three innexin-related proteins, called pannexins, have persisted in vertebrates, although it is not clear if they form intercellular channels (Panchin et al. 2000; Bruzzone et al. 2003). 7Å-resolution electron crystallographic structures of intercellular channels composed of either a carboxy-terminal truncation of Cx43 (Unger et al. 1999; Yeager and Harris 2007) or an M34A mutant of Cx26 (Oshima et al. 2007) are available. The overall pore morphologies are similar with the exception of a “plug” in the Cx26 channel pore. The density of this plug is substantively decreased by deletion of amino acids 2–7, suggesting that the amino-terminus contributes to this structure (Oshima et al. 2008). A 3.5-Å X-ray crystallographic structure has visualized the amino-terminus of Cx26 folded into the mouth of the channel without forming a plug, thought to be an image of the open channel conformation (Maeda et al. 2009). The amino-terminus has been physiologically implicated in voltage-gating of the Cx26 and Cx32 channels (Purnick et al. 2000; Oh et al. 2004), lending support to a role for the amino-terminus as a gating structure. However, Cx43 also shows voltage-gating, and its lack of any structure resembling a plug remains unresolved. A comparison of a 1985 intercellular channel structure (Makowski 1985) with the 2009 3.5Å structure (Maeda et al. 2009) summarizes a quarter-century of X-ray progress (Fig. 3).Open in a separate windowFigure 1.A diagram showing the multiple levels of gap junction structure. Individual connexins assemble intracellularly into hexamers, called connexons, which then traffic to the cell surface. There, they dock with connexons in an adjacent cell, assembling an axial channel spanning two plasma membranes and a narrow extracellular “gap.”Open in a separate windowFigure 2.Electron microscopy of gap junctions joining adjacent hepatocytes in the mouse. The gap junction (GJ) is seen as an area of close plasma membrane apposition, clearly distinct from the tight junction (TJ) joining these cells. (Inset A) A high magnification view of the gap junction revealing the 2–3 nm “gap” (white arrows) separating the plasma membranes. (Inset B) A freeze-fracture replica of a gap junction showing the characteristic particles on the protoplasmic (P) fracture face and pits on the ectoplasmic (E) fracture face. The particles and pits show considerable disorder in their packing with an average 9-nm center-to-center spacing.Open in a separate windowFigure 3.A comparison of axial sections through gap-junction structures deduced from X-ray diffraction. The 1985 data (Makowski 1985) were acquired from gap junctions isolated biochemically from mouse liver containing mixtures of Cx32 and Cx26. The intercellular channel (CHANNEL) is blocked at the two cytoplasmic surfaces by electron density at the channel mouths along the sixfold symmetry axis. The 2009 data (Maeda et al. 2009), acquired from three-dimensional crystals of recombinant Cx26, resolve this density at the channel opening as the amino-termini of the connexin proteins, the 2009 model possibly showing an open channel structure.Most cells express multiple connexins. These may co-oligomerize into the same (homomeric) or mixed (heteromeric) connexons, although only certain combinations are permitted (Falk et al. 1997; Segretain and Falk 2004). A connexon may dock with an identical connexon to form a homotypic intercellular channel or with a connexon containing different connexins to form a heterotypic channel (Dedek et al. 2006). Although only some assembly combinations are permitted (White et al. 1994), the number of possible different intercellular channels formed by this 21-member family is astonishingly large. This diversity has significance because intercellular channels composed of different connexins have different physiological properties, including single-channel conductances and multiple conductance states (Takens-Kwak and Jongsma 1992), as well as permeabilities to experimental tracers (Elfgang et al. 1995) and to biologically relevant permeants (Gaunt and Subak-Sharpe 1979; Veenstra et al. 1995; Bevans et al. 1998; Gong and Nicholson 2001; Goldberg et al. 2002; Ayad et al. 2006; Harris 2007).Opening of extrajunctional connexons in the plasma membrane, described as “hemichannel” activity, can be experimentally induced in a variety of cell types. Because first observations of hemichannel activity were in an oocyte expression system (Paul et al. 1991) and dissociated retinal horizontal cells (DeVries and Schwartz 1992), the possible functions of hemichannels composed of connexins and pannexins has enjoyed vigorous investigation (Goodenough and Paul 2003; Bennett et al. 2003; Locovei et al. 2006; Evans et al. 2006; Srinivas et al. 2007; Schenk et al. 2008; Thompson and MacVicar 2008; Anselmi et al. 2008; Goodenough and Paul 2003). Hemichannels have been implicated in various forms of paracrine signaling, for example in providing a pathway for extracellular release of ATP (Cotrina et al. 1998; Kang et al. 2008), glutamate (Ye et al. 2003), NAD⁺ (Bruzzone et al. 2000), and prostaglandins (Jiang and Cherian 2003).     

Cargo Recognition in Clathrin-Mediated Endocytosis

The endosomal system is expansive and complex, characterized by swift morphological transitions, dynamic remodeling of membrane constituents, and intracellular positioning changes. To properly navigate this ever-altering membrane labyrinth, transmembrane protein cargoes typically require specific sorting signals that are decoded by components of protein coats. The best-characterized sorting process within the endosomal system is the rapid internalization of select transmembrane proteins within clathrin-coated vesicles. Endocytic signals consist of linear motifs, conformational determinants, or covalent modifications in the cytosolic domains of transmembrane cargo. These signals are interpreted by a diverse set of clathrin-associated sorting proteins (CLASPs) that translocate from the cytosol to the inner face of the plasma membrane. Signal recognition by CLASPs is highly cooperative, involving additional interactions with phospholipids, Arf GTPases, other CLASPs, and clathrin, and is regulated by large conformational changes and covalent modifications. Related sorting events occur at other endosomal sorting stations.The internalization of a subset of plasma membrane proteins by clathrin-mediated endocytosis is one the best-characterized sorting processes that takes place in the endomembrane system of eukaryotic cells (Kirchhausen 2014). Selection of transmembrane proteins (referred to as “cargo”) for internalization by clathrin-mediated endocytosis involves recognition of endocytic signals in the cytosolic domains of the proteins by adaptors located in the inner layer of clathrin coats. Signal–adaptor interactions lead to concentration of the transmembrane proteins within clathrin-coated pits that eventually bud into the cytoplasm as clathrin-coated vesicles (Kirchhausen 2014). Transmembrane proteins that have endocytic signals are thus rapidly delivered to endosomes, whereas those that lack signals remain at the plasma membrane. This article summarizes recent progress in the elucidation of the mechanisms of signal recognition in clathrin-mediated endocytosis, with additional reference to related intracellular sorting events. Further information on this topic can be found in previous reviews (Bonifacino and Traub 2003; Traub 2009; Kelly and Owen 2011).