Retrograde Traffic from the Golgi to the Endoplasmic Reticulum

Proteins to be secreted are transported from the endoplasmic reticulum (ER) to the Golgi apparatus. The transport of these proteins requires the localization and activity of proteins that create ER exit sites, coat proteins to collect cargo and to reshape the membrane into a transport container, and address labels—SNARE proteins—to target the vesicles specifically to the Golgi apparatus. In addition some proteins may need export chaperones or export receptors to enable their exit into transport vesicles. ER export factors, SNAREs, and misfolded Golgi-resident proteins must all be retrieved from the Golgi to the ER again. This retrieval is also part of the organellar homeostasis pathway essential to maintaining the identity of the ER and of the Golgi apparatus. In this review, I will discuss the different processes in retrograde transport from the Golgi to the ER and highlight the mechanistic insights we have obtained in the last couple of years.Proteins that are exposed at the plasma membrane or populate a membrane-bounded organelle are synthesized into the endoplasmic reticulum (ER). In the ER, the folding of these proteins takes place and posttranslational modifications such as N-glycosylation and disulfide bridge formation occur. Upon adopting a suitable, often correct, conformation, proteins destined to locations beyond the ER are concentrated at so-called ER exit sites (ERES) and incorporated into nascent COPII-coated vesicles. These COPII vesicles eventually bud off the ER membrane and are transported to the Golgi (in yeast, Drosophila, and C. elegans) or the ER-Golgi intermediate compartment (in mammalian cells) (Schweizer et al. 1990; Kondylis and Rabouille 2003; Spang 2009; Witte et al. 2011).It is assumed that the vesicle coat is at least partially destabilized through the hydrolysis of GTP by the small GTPase Sar1 (Oka and Nakano 1994; Springer et al. 1999). However, some of the destabilized coat components have to stay on the vesicle until it has reached the Golgi apparatus because coat components participate in the recognition and the tethering process (Barlowe 1997; Cai et al. 2007; Lord et al. 2011; Zong et al. 2012). Subsequently, SNARE proteins on the vesicles (v-SNAREs) zipper up with cognate SNAREs on the Golgi (target SNAREs, t-SNAREs) to drive membrane fusion (Hay et al. 1998; Cao and Barlowe 2000; Parlati et al. 2002). The content of the ER-derived COPII vesicles is thereby released into the lumen of the cis-cisterna of the Golgi apparatus. Most proteins will continue their journey through the Golgi apparatus and encounter further modifications such as extension of the glycosylation tree or lipidation. However, some proteins, especially those involved in the fusion process, i.e., the v-SNAREs or proteins that act as export factors of the ER, such as Vma21, which is essential for export of the correctly folded and assembled V0 sector of the V-ATPase, need to be recycled back to the ER for another round of transport (Ballensiefen et al. 1998; Malkus et al. 2004). Moreover, cis-Golgi proteins are returned to the ER for quality/functional control (Todorow et al. 2000; Sato et al. 2004; Valkova et al. 2011). Finally, some ER-resident proteins, such as the ER Hsp70 chaperone BiP/Kar2, can escape the ER, but are captured at the cis-Golgi by the H/KDEL receptor Erd2 and returned to the ER (Lewis et al. 1990; Semenza et al. 1990; Aoe et al. 1997).Unfortunately, the retrograde transport route is also hijacked by toxins. For example, endocytosed cholera toxin subunit A contains a KDEL sequence and can thereby exploit the system to access the ER (Majoul et al. 1996, 1998). From there, it is retro-translocated into the cytoplasm where it can exert its detrimental function.     

Unconventional Functions for Clathrin,ESCRTs, and Other Endocytic Regulators in the Cytoskeleton,Cell Cycle,Nucleus, and Beyond: Links to Human Disease

The roles of clathrin, its regulators, and the ESCRT (endosomal sorting complex required for transport) proteins are well defined in endocytosis. These proteins can also participate in intracellular pathways that are independent of endocytosis and even independent of the membrane trafficking function of these proteins. These nonendocytic functions involve unconventional biochemical interactions for some endocytic regulators, but can also exploit known interactions for nonendocytic functions. The molecular basis for the involvement of endocytic regulators in unconventional functions that influence the cytoskeleton, cell cycle, signaling, and gene regulation are described here. Through these additional functions, endocytic regulators participate in pathways that affect infection, glucose metabolism, development, and cellular transformation, expanding their significance in human health and disease.The discovery and characterization of clathrin (Pearse 1975) initiated molecular definition of the many endocytosis regulators described in this collection, which mediate the clathrin-dependent and -independent pathways for membrane internalization (see Kirchhausen et al. 2014; Mayor et al. 2014; Merrifield and Kaksonen 2014). In accompanying reviews, we have seen how these endocytic pathways influence nutrition and metabolism (see Antonescu et al. 2014), signal transduction (see Bökel and Brand 2014; Di Fiore and von Zastrow 2014), neuronal function (see Morgan et al. 2013; Cosker and Segal 2014), infection and immunity (see ten Broeke et al. 2013; Cossart and Helenius 2014), tissue polarity and development (see Eaton and Martin-Belmonte 2014; Gonzalez-Gaitan and Jülicher 2014), and migration and metastasis (see Mellman and Yarden 2013). Recently, it has been established that some endocytic regulators have molecular properties that expand their functions beyond endocytosis. These include molecular interactions that affect the microtubule and actin cytoskeletons, nuclear translocation that influences gene regulation, and the formation of membrane-associated scaffolds that serve as signaling and sorting platforms. Through these diverse nonendocytic functions, endocytosis regulators play additional roles in cell division, pathogen infection, cell adhesion, and oncogenesis. In this article, we review the nonconventional behavior of endocytic regulators, first discussing the molecular properties that enable their moonlighting functions and then discussing the cellular processes and disease states that are influenced by these functions.     

Endocytosis,Signaling, and Beyond

The endocytic network comprises a vast and intricate system of membrane-delimited cell entry and cargo sorting routes running between biochemically and functionally distinct intracellular compartments. The endocytic network caters to the organization and redistribution of diverse subcellular components, and mediates appropriate shuttling and processing of materials acquired from neighboring cells or the extracellular milieu. Such trafficking logistics, despite their importance, represent only one facet of endocytic function. The endocytic network also plays a key role in organizing, mediating, and regulating cellular signal transduction events. Conversely, cellular signaling processes tightly control the endocytic pathway at different steps. The present article provides a perspective on the intimate relationships that exist between particular endocytic and cellular signaling processes in mammalian cells, within the context of understanding the impact of this nexus on integrated physiology.Molecular mechanisms governing the remarkable diversity of endocytic routes and trafficking steps are described elsewhere in the literature (see Bissig and Gruenberg 2013; Henne et al. 2013; Burd and Cullen 2014; Gautreau et al. 2014; Kirchhausen et al. 2014; Mayor et al. 2014; Merrifield and Kaksonen 2014; Piper et al. 2014). Moreover, these have been the focus of many studies in the last 30 years, and the topic has been covered by many excellent reviews, making it unnecessary for us to dwell on this aspect any further here (see, for instance, Howes et al. 2010; McMahon and Boucrot 2011; Sandvig et al. 2011; Parton and del Pozo 2013). Herein, we will instead concentrate our attention on how cellular regulatory mechanisms control endocytosis, as well as on how endocytic events impinge on cell functions. Emphasis will be placed, although not exclusively, on studies that analyze cellular networks using holistic approaches and in vivo analysis. Our aim is to give the reader a flavor of the deep embedding of endocytic processes within cellular programs, a concept we refer to as the endocytic matrix (Scita and Di Fiore 2010).     

The Endoplasmic Reticulum Chaperone BiP/GRP78 Is Important in the Structure and Function of the Human Cytomegalovirus Assembly Compartment

We previously demonstrated that the endoplasmic reticulum (ER) chaperone BiP functions in human cytomegalovirus (HCMV) assembly and egress. Here, we show that BiP localizes in two cytoplasmic structures in infected cells. Antibodies to the extreme C terminus, which includes BiP''s KDEL ER localization sequence, detect BiP in regions of condensed ER near the periphery of the cell. Antibodies to the full length, N terminus, or larger portion of the C terminus detect BiP in the assembly compartment. This inability of C-terminal antibodies to detect BiP in the assembly compartment suggests that BiP''s KDEL sequence is occluded in the assembly compartment. Depletion of BiP causes the condensed ER and assembly compartments to dissociate, indicating that BiP is important for their integrity. BiP and pp28 are in association in the assembly compartment, since antibodies that detect BiP in the assembly compartment coimmunoprecipitate pp28 and vice versa. In addition, BiP and pp28 copurify with other assembly compartment components on sucrose gradients. BiP also coimmunoprecipitates TRS1. Previous data show that cells infected with a TRS1-deficient virus have cytoplasmic and assembly compartment defects like those seen when BiP is depleted. We show that a fraction of TRS1 purifies with the assembly compartment. These findings suggest that BiP and TRS1 share a function in assembly compartment maintenance. In summary, BiP is diverted from the ER to associate with pp28 and TRS1, contributing to the integrity and function of the assembly compartment.Human cytomegalovirus (HCMV), the largest of the human herpesviruses, is capable of encoding over 200 proteins, which are expressed in temporal fashion as immediate-early, early, delayed-early, and late genes. Despite the extensive coding capacity of HCMV, its replication cycle is slow. During this protracted period, the virus must maintain optimal replication conditions in the host cell. However, the increasing strain of the infection induces cellular stress responses with consequences that may be deleterious to the progress of the infection. We and others have previously shown that HCMV has multiple mechanisms to deal with the deleterious aspects of cellular stress responses while maintaining beneficial ones (2, 8-10, 14, 17, 18, 22-24, 26, 27, 50, 51).An example of these mechanisms is the viral control of endoplasmic reticulum (ER) stress and the unfolded protein response (UPR). Due to the number of HCMV proteins that are glycosylated, or receive other ER-dependent posttranslational modifications, the load of proteins in the ER can exceed its capacity, resulting in ER stress and the activation of the UPR (18, 47, 51). However, we and others have shown that HCMV controls and modulates the UPR, maintaining aspects that may benefit the viral infection while inhibiting aspects that would be detrimental (18, 51).The UPR is normally controlled by transmembrane sensors which initiate the complex UPR signaling cascade when activated by ER stress (reviewed in references 20, 35, 38, and 52). The ER molecular chaperone BiP (immunoglobulin heavy chain-binding protein), also called glucose-regulated protein 78 (GRP78), is believed to bind these sensors and keep them inactive during unstressed conditions. However, when unfolded or misfolded proteins accumulate in the ER, BiP leaves these sensors to perform its chaperone function, thus allowing the sensors to activate UPR signaling. We have previously shown that during HCMV infection, BiP is vastly overproduced (8), suggesting that BiP may have other functions in the viral infection. Indeed, it has been shown that BiP binds to the viral proteins US2 and US11; this interaction is necessary for the virus-mediated degradation of major histocompatibility complex class I and II (15, 47). Further, we have shown that depletion of BiP, using either the BiP-specific subtilase cytotoxin SubAB (32) or short hairpin RNAs, caused infectious virion formation in the cytoplasm to cease and nucleocapsids to accumulate just outside the outer nuclear membrane (8). This result suggested that BiP has a significant role in virion formation and cytoplasmic egress.Although the exact mechanism of virion formation in the cytoplasm is not well understood, studies have identified a perinuclear structure, referred to as the cytoplasmic assembly compartment, that is involved in the process. Several viral proteins, for example, tegument proteins (pp28, pp65) (36) and viral glycoproteins (gB, gH, gL, gO, gp65) (36, 46), have been identified as part of this structure. Defining the exact origin of this compartment has been complicated by the observation of specific organellar markers in and around the compartment, while other markers of the same organelles are not detected. For example, immunofluorescence examination suggests that the early endosomal marker early endosome antigen 1 (EEA1) has been observed in the center of the assembly compartment (12, 13); however, Rab4 and Rab5, other early endosomal markers, were not detected (16). Such observations suggest that the virus directs specific viral and cellular proteins to the assembly compartment as needed for assembly compartment function.In the present study, we further examine the role of BiP during an HCMV infection, including its localization and interactions with other proteins. We show here that in infected cells, BiP localizes in two distinct structures, regions of condensed ER near the periphery of the cell and the assembly compartment. The data suggest that BiP diversion from the ER to the assembly compartment is due to occlusion of its ER localization signal. Depletion of BiP causes both condensed ER and assembly compartments to disperse, indicating that BiP is important for their formation or maintenance. BiP and pp28 appear to associate in the assembly compartment, since BiP from the assembly compartment coimmunoprecipitates pp28 and vice versa. In addition, both BiP and pp28 copurify with the assembly compartment on sucrose gradients. BiP also coimmunoprecipitates TRS1. Previous studies (1, 4) have shown that cells infected with HCMV with a mutation in the TRS1 gene show cytoplasmic and assembly compartment defects like those seen when BiP is depleted (reference 8 and the studies presented below). We show that a fraction of TRS1 purifies with the assembly compartment, indicating a shared assembly compartment function with BiP. In summary, our data suggest that BiP is diverted from the ER to associate with pp28 and TRS1, contributing to the integrity and function of the assembly compartment.     

Sequential Recruitment of the Endoplasmic Reticulum and Chloroplasts for Plant Potyvirus Replication

The replication of positive-strand RNA viruses occurs in cytoplasmic membrane-bound virus replication complexes (VRCs). Depending on the virus, distinct cellular organelles such as the endoplasmic reticulum (ER), chloroplast, mitochondrion, endosome, and peroxisome are recruited for the formation of VRC-associated membranous structures. Previously, the 6,000-molecular-weight protein (6K) of plant potyviruses was shown to be an integral membrane protein that induces the formation of 6K-containing membranous vesicles at endoplasmic reticulum (ER) exit sites for potyvirus genome replication. Here, we present evidence that the 6K-induced vesicles predominantly target chloroplasts, where they amalgamate and induce chloroplast membrane invaginations. The vesicular transport pathway and actomyosin motility system are involved in the trafficking of the 6K vesicles from the ER to chloroplasts. Viral RNA, double-stranded RNA, and viral replicase components are concentrated at the 6K vesicles that associate with chloroplasts in infected cells, suggesting that these chloroplast-bound 6K vesicles are the site for potyvirus replication. Taken together, these results suggest that plant potyviruses sequentially recruit the ER and chloroplasts for their genome replication.The replication of eukaryotic positive-strand RNA viruses in infected cells is closely associated with unique virus-induced intracellular membranous vesicles (22). These membranous vesicles have been proposed to provide a scaffold for anchoring the virus replication complex (VRC), confine the process of RNA replication to a specific safeguarded cytoplasmic location, and prevent the activation of certain host defense mechanisms that can be triggered by double-stranded RNA (dsRNA) intermediates during virus replication (33, 47). Depending on the type of virus, the virus-induced membranous vesicles are derived from various intracellular organelles in the host. Many plant and animal viruses remodel and utilize the endoplasmic reticulum (ER) in VRCs (1, 6, 17, 33, 34, 36, 38, 39, 46). Other cellular organelles such as endosomes, lysomes, chloroplasts, peroxisomes, and mitochondria have also been suggest to be the replication site for togaviruses, tymoviruses, and tombusviruses, respectively (25, 27, 31). Given that the ER appears to be the site where the host cell translation machinery is hijacked for the biosynthesis of the first set of viral proteins, the subcellular location of virus replication (either in the vicinity of the ER or elsewhere) and the mechanism of transport to locations other than the ER are poorly understood.Plant potyviruses, accounting for ∼30% of known plant viruses including many agriculturally important viruses, e.g., Turnip mosaic virus (TuMV), Maize dwarf mosaic virus (MDMV), Tobacco etch virus (TEV), and Potato virus Y (PVY), are related to picornaviruses and picorna-like viruses (20, 21, 43). The potyviral genome is a single-stranded positive-sense RNA of about 10 kb in length and encodes at least 11 mature viral proteins (8, 43). Of these 11 proteins, the 6-kDa protein (designated 6K or 6K2) contains a central hydrophobic domain (35). In seminal work, Carrington and colleagues determined that 6K induces the formation of the ER-derived vesicles for TEV replication (35, 38). More recently, viral proteins required for replication and several host factors, namely, eukaryotic initiation factor (isoform) 4E, poly(A)-binding protein, eukaryotic elongation factor 1A, and heat shock cognate 70-3 protein, have been shown to associate with the TuMV 6K-induced vesicles (9, 41), raising the possibility that the potyviral 6K vesicles represent sites of viral genome replication. Furthermore, we have demonstrated that the biogenesis of the potyviral 6K vesicles occurs at COPII-accumulating ER exit sites (ERES) on the ER membrane (45). In this study, we further studied the trafficking of 6K-induced vesicles and found that the 6K-induced mobile vesicles trafficked predominantly from the ER to the periphery of chloroplasts. We show that these 6K vesicles docked on the outer chloroplast envelope and induced chloroplast invaginations. The chloroplast-associated 6K vesicles contained viral replicase components and dsRNA and were concentrated with viral RNA. We provide evidence that the early secretory pathway and actomyosin motility system were required for the trafficking of 6K vesicles from the ER to chloroplasts. These results suggest that plant potyviruses sequentially recruit the ER and chloroplasts for their genome replication.     

Endocytosis of Viruses and Bacteria

Of the many pathogens that infect humans and animals, a large number use cells of the host organism as protected sites for replication. To reach the relevant intracellular compartments, they take advantage of the endocytosis machinery and exploit the network of endocytic organelles for penetration into the cytosol or as sites of replication. In this review, we discuss the endocytic entry processes used by viruses and bacteria and compare the strategies used by these dissimilar classes of pathogens.Many of the most widespread and devastating diseases in humans and livestock are caused by viruses and bacteria that enter cells for replication. Being obligate intracellular parasites, viruses have no choice. They must transport their genome to the cytosol or nucleus of infected cells to multiply and generate progeny. Bacteria and eukaryotic parasites do have other options; most of them can replicate on their own. However, some have evolved to take advantage of the protected environment in the cytosol or in cytoplasmic vacuoles of animal cells as a niche favorable for growth and multiplication. In both cases (viruses and intracellular bacteria), the outcome is often destructive for the host cell and host organism. The mortality and morbidity caused by infectious diseases worldwide provide a strong rationale for research into pathogen–host cell interactions and for pursuing the detailed mechanisms of transmission and dissemination. The study of viruses and bacteria can, moreover, provide invaluable insights into fundamental aspects of cell biology.Here, we focus on the mechanisms by which viral and bacterial pathogens exploit the endocytosis machinery for host cell entry and replication. Among recent reviews on this topic, dedicated uniquely to either mammalian viruses or bacterial pathogens, we recommend the following: Cossart and Sansonetti (2004); Pizarro-Cerda and Cossart (2006); Kumar and Valdivia (2009); Cossart and Roy (2010); Mercer et al. (2010b); Grove and Marsh (2011); Kubo et al. (2012); Vazquez-Calvo et al. (2012a); Sun et al. (2013).The term “endocytosis” is used herein in its widest sense, that is, to cover all processes whereby fluid, solutes, ligands, and components of the plasma membrane as well as particles (including pathogenic agents) are internalized by cells through the invagination of the plasma membrane and the scission of membrane vesicles or vacuoles. This differs from current practice in the bacterial pathogenesis field, where the term “endocytosis” is generally reserved for the internalization of molecules or small objects, whereas the uptake of bacteria into nonprofessional phagocytes is called “internalization” or “bacterial-induced phagocytosis.” In addition, the term “phagocytosis” is reserved for internalization of bacteria by professional phagocytes (macrophages, polymorphonuclear leucocytes, dendritic cells, and amoebae), a process that generally but not always leads to the destruction of the ingested bacteria (Swanson et al. 1999; May and Machesky 2001; Henry et al. 2004; Zhang et al. 2010). With a few exceptions, we will not discuss phagocytosis of bacteria or the endocytosis of protozoan parasites such as Toxoplasma and Plasmodium (Robibaro et al. 2001).     

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).     

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.     

Fibronectins,Their Fibrillogenesis,and In Vivo Functions

Fibronectin (FN) is a multidomain protein with the ability to bind simultaneously to cell surface receptors, collagen, proteoglycans, and other FN molecules. Many of these domains and interactions are also involved in the assembly of FN dimers into a multimeric fibrillar matrix. When, where, and how FN binds to its various partners must be controlled and coordinated during fibrillogenesis. Steps in the process of FN fibrillogenesis including FN self-association, receptor activities, and intracellular pathways have been under intense investigation for years. In this review, the domain organization of FN including the extra domains and variable region that are controlled by alternative splicing are described. We discuss how FN–FN and cell–FN interactions play essential roles in the initiation and progression of matrix assembly using complementary results from cell culture and embryonic model systems that have enhanced our understanding of this process.As a ubiquitous component of the extracellular matrix (ECM), fibronectin (FN) provides essential connections to cells through integrins and other receptors and regulates cell adhesion, migration, and differentiation. FN is secreted as a large dimeric glycoprotein with subunits that range in size from 230 kDa to 270 kDa (Mosher 1989; Hynes 1990). Variation in subunit size depends primarily on alternative splicing. FN was first isolated from blood more than 60 years ago (Edsall 1978), and this form is called plasma FN. The other major form, called cellular FN, is abundant in the fibrillar matrices of most tissues. Although FN is probably best known for promoting attachment of cells to surfaces, this multidomain protein has many interesting structural features and functional roles beyond cell adhesion.FN is composed of three different types of modules termed type I, II, and III repeats (Fig. 1) (Petersen et al. 1983; Hynes 1990). These repeats have distinct structures. Although the conformations of type I and type II repeats are maintained by pairs of intramodule disulfide bonds, the type III repeat is a 7-stranded β-barrel structure that lacks disulfide bonds (Main et al. 1992; Leahy et al. 1996, 1992) and, therefore, can undergo conformational changes. FN type III repeats are widely distributed among animal, bacterial, and plant proteins and are found in both extracellular and intracellular proteins (Bork and Doolittle 1992; Tsyguelnaia and Doolittle 1998).Open in a separate windowFigure 1.FN domain organization and isoforms. Each FN monomer has a modular structure consisting of 12 type I repeats (cylinders), 2 type II repeats (diamonds), and 15 constitutive type III repeats (hexagons). Two additional type III repeats (EIIIA and EIIIB, green) are included or omitted by alternative splicing. The third region of alternative splicing, the V region (green box), is included (V120), excluded (V0), or partially included (V95, V64, V89). Sets of modules comprise domains for binding to other extracellular molecules as indicated. Domains required for fibrillogenesis are in red: the assembly domain (repeats I_1-5) binds FN, III_9-10 contains the RGD and synergy sequences for integrin binding, and the carboxy-terminal cysteines form the disulfide-bonded FN dimer (‖). The III_1-2 domain (light red) has two FN binding sites that are important for fibrillogenesis. The amino-terminal 70-kDa fragment contains assembly and gelatin-binding domains and is routinely used in FN binding and matrix assembly studies.Sets of adjacent modules form binding domains for a variety of proteins and carbohydrates (Fig. 1). ECM proteins, including FN, bind to cells via integrin receptors, αβ heterodimers with two transmembrane subunits (Hynes 2002). FN-binding integrins have specificity for one of the two cell-binding sites within FN, either the RGD-dependent cell-binding domain in III₁₀ (Pierschbacher and Ruoslahti 1984) or the CS1 segment of the alternatively spliced V region (IIICS) (Wayner et al. 1989; Guan and Hynes 1990). Some integrins require a synergy sequence in repeat III₉ for maximal interactions with FN (Aota et al. 1994; Bowditch et al. 1994). Another family of cell surface receptors is the syndecans, single-chain transmembrane proteoglycans (Couchman 2010). Syndecans use their glycosaminoglycan (GAG) chains to interact with FN at its carboxy-terminal heparin-binding (HepII) domain (Fig. 1) (Saunders and Bernfield 1988; Woods et al. 2000), which binds to heparin, heparan sulfate, and chondroitin sulfate GAGs (Hynes 1990; Barkalow and Schwarzbauer 1994). Syndecan binding to the HepII domain enhances integrin-mediated cell spreading and intracellular signaling, suggesting that syndecans act as coreceptors with integrins in cell–FN binding (Woods and Couchman 1998; Morgan et al. 2007).A major site for FN self-association is within the amino-terminal assembly domain spanning the first five type I repeats (I_1-5) (Fig. 1) (McKeown-Longo and Mosher 1985; McDonald et al. 1987; Schwarzbauer 1991b; Sottile et al. 1991). This domain plays an essential role in FN fibrillogenesis. As a major blood protein, FN interacts with fibrin during blood coagulation, also using the I_1-5 domain (Mosher 1989; Hynes 1990). As fibrin polymerizes, factor XIII transglutaminase covalently cross-links glutamine residues near the amino terminus of FN to fibrin α chains (Mosher 1975; Corbett et al. 1997). The amino-terminal domain has multiple binding partners in addition to FN and fibrin; these include heparin, S. aureus, and other bacteria, thrombospondin-1, and tenascin-C (Hynes 1990; Ingham et al. 2004; Schwarz-Linek et al. 2006). Adjacent to this domain is the gelatin/collagen-binding domain composed of type I and type II modules (Ingham et al. 1988). This domain also binds to tissue transglutaminase (Radek et al. 1993) and fibrillin-1 (Sabatier et al. 2009). Within the 15 type III repeats reside several FN binding sites that interact with the amino-terminal assembly domain as well as three sites of alternative splicing that generate multiple isoforms. At the carboxyl terminus is a pair of cysteine residues that form the FN dimer through antiparallel disulfide bonds (Hynes 1990). This dimerization may be facilitated by disulfide isomerase activity located in the last set of type I repeats (Langenbach and Sottile 1999).The diverse set of binding domains provides FN with the ability to interact simultaneously with other FN molecules, other ECM components (e.g., collagens and proteoglycans), cell surface receptors, and extracellular enzymes (Pankov and Yamada 2002; Fogelgren et al. 2005; Hynes 2009; Singh et al. 2010). Multitasking by FN probably underlies its essential role during embryogenesis (George et al. 1993). Furthermore, FN''s interactions can be modulated by exposure or sequestration of its binding sites within matrix fibrils, through the presence of ECM proteins that bind to FN, or through variation in structure by alternative splicing.     

Dormant Origins,the Licensing Checkpoint,and the Response to Replicative Stresses

Only ∼10% of replication origins that are licensed by loading minichromosome maintenance 2-7 (MCM2-7) complexes are normally used, with the majority remaining dormant. If replication fork progression is inhibited, nearby dormant origins initiate to ensure that all of the chromosomal DNA is replicated. At the same time, DNA damage-response kinases are activated, which preferentially suppress the assembly of new replication factories. This diverts initiation events away from completely new areas of the genome toward regions experiencing replicative stress. Mice hypomorphic for MCM2-7, which activate fewer dormant origins in response to replication inhibition, are cancer-prone and are genetically unstable. The licensing checkpoint delays entry into S phase if an insufficient number of origins have been licensed. In contrast, humans with Meier-Gorlin syndrome have mutations in pre-RC proteins and show defects in cell proliferation that may be a consequence of chronic activation of the licensing checkpoint.Replicating the large amount of DNA in eukaryotic cells is a complex task, requiring the activation of hundreds or thousands of origins spread throughout the genome. To maintain genetic stability, it is essential that during S phase genomic DNA is precisely duplicated, with no sections of DNA left unreplicated and no section of DNA replicated more than once. To prevent re-replication, cells divide the process of DNA replication into two non-overlapping phases. Prior to S phase, origins are licensed by the binding of minichromosome maintenance 2-7 (MCM2-7) double hexamers (Gillespie et al. 2001; Blow and Dutta 2005; Arias and Walter 2007). During S phase, these are activated as the core of the CMG (Cdc45-MCM-GINS) replicative helicase (Moyer et al. 2006; Ilves et al. 2010). Prior to the onset of S phase, licensing proteins are down-regulated or inhibited, so that no more origins can be licensed (Wohlschlegel et al. 2000; Tada et al. 2001; Li et al. 2003; Li and Blow 2005). One consequence of using this mechanism for preventing re-replication of DNA is that it is imperative that enough origins are licensed prior to S-phase entry, so that no regions of the genome remain unreplicated, even if some replication forks stall or some origins fail to initiate (Blow et al. 2011). Metazoan cells employ a licensing checkpoint to monitor that sufficient origins are licensed, inhibiting S-phase entry until this is established (Shreeram et al. 2002; Blow and Gillespie 2008).Here we review recent research showing how cells ensure complete genome duplication by licensing more replication origins in G1 than are normally used during S phase. The otherwise dormant replication origins become important for ensuring the completion of DNA replication if replication forks stall or are inhibited during S phase. We also review research showing how the licensing checkpoint ensures that a large enough number of origins are licensed before cells embark on S phase.     

Nonvesicular Lipid Transfer from the Endoplasmic Reticulum

The transport of lipids from their synthesis site at the endoplasmic reticulum (ER) to different target membranes could be mediated by both vesicular and nonvesicular transport mechanisms. Nonvesicular lipid transport appears to be the major transport route of certain lipid species, and could be mediated by either spontaneous lipid transport or by lipid-transfer proteins (LTPs). Although nonvesicular lipid transport has been extensively studied for more than four decades, its underlying mechanism, advantage and regulation, have not been fully explored. In particular, the function of LTPs and their involvement in intracellular lipid movement remain largely controversial. In this article, we describe the pathways by which lipids are synthesized at the ER and delivered to different cellular membranes, and discuss the role of LTPs in lipid transport both in vitro and in intact cells.The endoplasmic reticulum (ER) is a large interconnected membrane network that plays a major role in lipid biosynthesis in eukaryotic cells (Borgese et al. 2006). Newly synthesized lipids at the ER are then delivered to different cellular membranes or organelles, each of which shows unique lipid and protein composition and executes distinct cellular function (Holthuis et al. 2003). The transport of lipids from the ER can be mediated by both vesicular and nonvesicular transport mechanisms. Vesicular transport, as opposed to nonvesicular lipid transport, requires metabolic energy, intact cytoskeleton, and connection to the vesicular transport machinery (Kaplan and Simoni 1985a; Voelker 1990; Vance et al. 1991). Although vesicular lipid transport mediates the bulk transport of many lipids, increasing lines of evidence suggest that nonvesicular lipid transport is the major transport route for certain lipid types (Lev 2010). Nonvesicular lipid transport between membranes could be mediated by spontaneous lipid transport, in which a lipid monomer is diffused through the cytosol from a donor to the acceptor membrane. Given that most cellular lipids are highly hydrophobic, their diffusion through an aqueous phase is very slow and insufficient to support substantial transport of most lipids (Jones and Thompson 1989; Mesmin and Maxfield 2009). Nevertheless, spontaneous lipid transport can be greatly facilitated at membrane contact sites (MCSs) (Levine 2004; Holthuis and Levine 2005) and/or by lipid-transfer proteins (LTPs) (Lev 2010). MCSs are defined as small cytosolic gaps of 10–20 nm between the ER membranes and virtually all cellular organelles (Levine 2004; Lebiedzinska et al. 2009), whereas LTPs are intracellular proteins that can carry a lipid monomer in a hydrophobic pocket and transfer it between membranes through an aqueous phase. LTPs were initially discovered as soluble factors that accelerate the exchange or net transfer of different lipid species between membranes in vitro (Wirtz and Zilversmit 1968). Subsequently, many LTPs have been isolated, cloned, and crystallized. LTPs have been identified in all eukaryotes, in plants, and in bacteria, and according to their sequence and structure similarity have been subdivided into different protein families including SEC14, PITP (phosphatidylinositol-transfer protein), START (StAR-related lipid transfer), GLTP (glycolipid transfer protein), SCP-2 (nonspecific LTPs), and OSBP (oxysterol-binding protein)/ORP (OSBP-related proteins) (D’Angelo et al. 2008). In general, LTPs show specificity for one or more lipid types, and may contain only a single lipid-transfer domain (LTD), or additional structural domains with varying functions (Lev 2010). Over the past 40 years, LTPs have been extensively studied and the major principles of their action mode have been established from both biophysical measurements in vitro and structural data (Lev 2010). Nevertheless, the precise function of LTPs in intact cells remains controversial and a subject of an active field of research. In this article, we briefly describe how lipids are synthesized in the ER and delivered to different target membranes, and discuss how LTPs influence lipid transport in vitro and in intact cells.     

Molecular Mechanisms of Fibroblast Growth Factor Signaling in Physiology and Pathology

Fibroblast growth factors (FGFs) signal in a paracrine or endocrine fashion to mediate a myriad of biological activities, ranging from issuing developmental cues, maintaining tissue homeostasis, and regulating metabolic processes. FGFs carry out their diverse functions by binding and dimerizing FGF receptors (FGFRs) in a heparan sulfate (HS) cofactor- or Klotho coreceptor-assisted manner. The accumulated wealth of structural and biophysical data in the past decade has transformed our understanding of the mechanism of FGF signaling in human health and development, and has provided novel concepts in receptor tyrosine kinase (RTK) signaling. Among these contributions are the elucidation of HS-assisted receptor dimerization, delineation of the molecular determinants of ligand–receptor specificity, tyrosine kinase regulation, receptor cis-autoinhibition, and tyrosine trans-autophosphorylation. These structural studies have also revealed how disease-associated mutations highjack the physiological mechanisms of FGFR regulation to contribute to human diseases. In this paper, we will discuss the structurally and biophysically derived mechanisms of FGF signaling, and how the insights gained may guide the development of therapies for treatment of a diverse array of human diseases.Fibroblast growth factor (FGF) signaling fulfills essential roles in metazoan development and metabolism. A wealth of literature has documented the requirement for FGF signaling in multiple processes during embryogenesis, including implantation (Feldman et al. 1995), gastrulation (Sun et al. 1999), somitogenesis (Dubrulle and Pourquie 2004; Wahl et al. 2007; Lee et al. 2009; Naiche et al. 2011; Niwa et al. 2011), body plan formation (Martin 1998; Rodriguez Esteban et al. 1999; Tanaka et al. 2005; Mariani et al. 2008), morphogenesis (Metzger et al. 2008; Makarenkova et al. 2009), and organogenesis (Goldfarb 1996; Kato and Sekine 1999; Sekine et al. 1999; Sun et al. 1999; Colvin et al. 2001; Serls et al. 2005; Vega-Hernandez et al. 2011). Recent clinical and biochemical data have uncovered unexpected roles for FGF signaling in metabolic processes, including phosphate/vitamin D homeostasis (Consortium 2000; Razzaque and Lanske 2007; Nakatani et al. 2009; Gattineni et al. 2011; Kir et al. 2011), cholesterol/bile acid homeostasis (Yu et al. 2000a; Holt et al. 2003), and glucose/lipid metabolism (Fu et al. 2004; Moyers et al. 2007). Highlighting its diverse biology, deranged FGF signaling contributes to many human diseases, such as congenital craniosynostosis and dwarfism syndromes (Naski et al. 1996; Wilkie et al. 2002, 2005), Kallmann syndrome (Dode et al. 2003; Pitteloud et al. 2006a), hearing loss (Tekin et al. 2007, 2008), and renal phosphate wasting disorders (Shimada et al. 2001; White et al. 2001), as well as many acquired forms of cancers (Rand et al. 2005; Pollock et al. 2007; Gartside et al. 2009; di Martino et al. 2012). Endocrine FGFs have also been implicated in the progression of acquired metabolic disorders, including chronic kidney disease (Fliser et al. 2007), obesity (Inagaki et al. 2007; Moyers et al. 2007; Reinehr et al. 2012), and insulin resistance (Fu et al. 2004; Chen et al. 2008b; Chateau et al. 2010; Huang et al. 2011), giving rise to many opportunities for drug discovery in the field of FGF biology (Beenken and Mohammadi 2012).Based on sequence homology and phylogeny, the 18 mammalian FGFs are grouped into six subfamilies (Ornitz and Itoh 2001; Popovici et al. 2005; Itoh and Ornitz 2011). Five of these subfamilies act in a paracrine fashion, namely, the FGF1 subfamily (FGF1 and FGF2), the FGF4 subfamily (FGF4, FGF5, and FGF6), the FGF7 subfamily (FGF3, FGF7, FGF10, and FGF22), the FGF8 subfamily (FGF8, FGF17, and FGF18), and the FGF9 subfamily (FGF9, FGF16, and FGF20). In contrast, the FGF19 subfamily (FGF19, FGF21, and FGF23) signals in an endocrine manner (Beenken and Mohammadi 2012). FGFs exert their pleiotropic effects by binding and activating the FGF receptor (FGFR) subfamily of receptor tyrosine kinases that are coded by four genes (FGFR1, FGFR2, FGFR3, and FGFR4) in mammals (Johnson and Williams 1993; Mohammadi et al. 2005b). The extracellular domain of FGFRs consists of three immunoglobulin (Ig)-like domains (D1, D2, and D3), and the intracellular domain harbors the conserved tyrosine kinase domain flanked by the flexible amino-terminal juxtamembrane linker and carboxy-terminal tail (Lee et al. 1989; Dionne et al. 1991; Givol and Yayon 1992). A unique feature of FGFRs is the presence of a contiguous segment of glutamic and aspartic acids in the D1–D2 linker, termed the acid box (AB). The two-membrane proximal D2 and D3 and the intervening D2–D3 linker are necessary and sufficient for ligand binding/specificity (Dionne et al. 1990; Johnson et al. 1990), whereas D1 and the D1–D2 linker are implicated in receptor autoinhibition (Wang et al. 1995; Roghani and Moscatelli 2007; Kalinina et al. 2012). Alternative splicing and translational initiation further diversify both ligands and receptors. The amino-terminal regions of FGF8 and FGF17 can be differentially spliced to yield FGF8a, FGF8b, FGF8e, FGF8f (Gemel et al. 1996; Blunt et al. 1997), and FGF17a and FGF17b isoforms (Xu et al. 1999), whereas cytosine-thymine-guanine (CTG)-mediated translational initiation gives rise to multiple high molecular weight isoforms of FGF2 and FGF3 (Florkiewicz and Sommer 1989; Prats et al. 1989; Acland et al. 1990). The tissue-specific alternative splicing in D3 of FGFR1, FGFR2, and FGFR3 yields “b” and “c” receptor isoforms which, along with their temporal and spatial expression patterns, is the major regulator of FGF–FGFR specificity/promiscuity (Orr-Urtreger et al. 1993; Ornitz et al. 1996; Zhang et al. 2006). A large body of structural data on FGF–FGFR complexes has begun to reveal the intricate mechanisms by which different FGFs and FGFRs combine selectively to generate quantitatively and qualitatively different intracellular signals, culminating in distinct biological responses. In addition, these structural data have unveiled how pathogenic mutations hijack the normal physiological mechanisms of FGFR regulation to lead to pathogenesis. We will discuss the current state of the structural biology of the FGF–FGFR system, lessons learned from studying the mechanism of action of pathogenic mutations, and how the structural data are beginning to shape and advance the translational research.     

Origin of Microglia: Current Concepts and Past Controversies

Microglia are the resident macrophages of the central nervous system (CNS), which sit in close proximity to neural structures and are intimately involved in brain homeostasis. The microglial population also plays fundamental roles during neuronal expansion and differentiation, as well as in the perinatal establishment of synaptic circuits. Any change in the normal brain environment results in microglial activation, which can be detrimental if not appropriately regulated. Aberrant microglial function has been linked to the development of several neurological and psychiatric diseases. However, microglia also possess potent immunoregulatory and regenerative capacities, making them attractive targets for therapeutic manipulation. Such rationale manipulations will, however, require in-depth knowledge of their origins and the molecular mechanisms underlying their homeostasis. Here, we discuss the latest advances in our understanding of the origin, differentiation, and homeostasis of microglial cells and their myelomonocytic relatives in the CNS.Microglia are the resident macrophages of the central nervous system (CNS), which are uniformly distributed throughout the brain and spinal cord with increased densities in neuronal nuclei, including the Substantia nigra in the midbrain (Lawson et al. 1990; Perry 1998). They belong to the nonneuronal glial cell compartment and their function is crucial to maintenance of the CNS in both health and disease (Ransohoff and Perry 2009; Perry et al. 2010; Ransohoff and Cardona 2010; Prinz and Priller 2014).Two key functional features define microglia: immune defense and maintenance of CNS homeostasis. As part of the innate immune system, microglia constantly sample their environment, scanning and surveying for signals of external danger (Davalos et al. 2005; Nimmerjahn et al. 2005; Lehnardt 2010), such as those from invading pathogens, or internal danger signals generated locally by damaged or dying cells (Bessis et al. 2007; Hanisch and Kettenmann 2007). Detection of such signals initiates a program of microglial responses that aim to resolve the injury, protect the CNS from the effects of the inflammation, and support tissue repair and remodeling (Minghetti and Levi 1998; Goldmann and Prinz 2013).Microglia are also emerging as crucial contributors to brain homeostasis through control of neuronal proliferation and differentiation, as well as influencing formation of synaptic connections (Lawson et al. 1990; Perry 1998; Hughes 2012; Blank and Prinz 2013). Recent imaging studies revealed dynamic interactions between microglia and synaptic connections in the healthy brain, which contributed to the modification and elimination of synaptic structures (Perry et al. 2010; Tremblay et al. 2010; Bialas and Stevens 2013). In the prenatal brain, microglia regulate the wiring of forebrain circuits, controlling the growth of dopaminergic axons in the forebrain and the laminar positioning of subsets of neocortical interneurons (Squarzoni et al. 2014). In the postnatal brain, microglia-mediated synaptic pruning is similarly required for the remodeling of neural circuits (Paolicelli et al. 2011; Schafer et al. 2012). In summary, microglia occupy a central position in defense and maintenance of the CNS and, as a consequence, are a key target for the treatment of neurological and psychiatric disorders.Although microglia have been studied for decades, a long history of experimental misinterpretation meant that their true origins remained debated until recently. Although we knew that microglial progenitors invaded the brain rudiment at very early stages of embryonic development (Alliot et al. 1999; Ransohoff and Perry 2009), it has now been established that microglia arise from yolk sac (YS)-primitive macrophages, which persist in the CNS into adulthood (Davalos et al. 2005; Nimmerjahn et al. 2005; Ginhoux et al. 2010, 2013; Kierdorf and Prinz 2013; Kierdorf et al. 2013a). Moreover, early embryonic brain colonization by microglia is conserved across vertebrate species, implying that it is essential for early brain development (Herbomel et al. 2001; Bessis et al. 2007; Hanisch and Kettenmann 2007; Verney et al. 2010; Schlegelmilch et al. 2011; Swinnen et al. 2013). In this review, we will present the latest findings in the field of microglial ontogeny, which provide new insights into their roles in health and disease.     

What Was the Real Contribution of Endosymbionts to the Eukaryotic Nucleus? Insights from Photosynthetic Eukaryotes

Eukaryotic genomes are composed of genes of different evolutionary origins. This is especially true in the case of photosynthetic eukaryotes, which, in addition to typical eukaryotic genes and genes of mitochondrial origin, also contain genes coming from the primary plastids and, in the case of secondary photosynthetic eukaryotes, many genes provided by the nuclei of red or green algal endosymbionts. Phylogenomic analyses have been applied to detect those genes and, in some cases, have led to proposing the existence of cryptic, no longer visible endosymbionts. However, detecting them is a very difficult task because, most often, those genes were acquired a long time ago and their phylogenetic signal has been heavily erased. We revisit here two examples, the putative cryptic endosymbiosis of green algae in diatoms and chromerids and of Chlamydiae in the first photosynthetic eukaryotes. We show that the evidence sustaining them has been largely overestimated, and we insist on the necessity of careful, accurate phylogenetic analyses to obtain reliable results.Today it is widely accepted that photosynthesis originated in eukaryotes by the endosymbiosis of a cyanobacterium within a heterotrophic eukaryotic host. This occurred in a lineage that subsequently diversified to give rise to the three contemporary groups of primary photosynthetic eukaryotes: Viridiplantae (including green algae and land plants), Rhodophyta and Glaucophyta, grouped collectively within a unique eukaryotic superphylum called Archaeplastida (Adl et al. 2005) or Plantae (Cavalier-Smith 1982). Recently, a second case of primary endosymbioses has been unveiled thanks to the characterization of Paulinella chromatophora, a filose amoeba that hosts a cyanobacterium with a reduced genome that has been described as “a plastid in the making” (Marin et al. 2005; Keeling and Archibald 2008; Nowack et al. 2008). Primary endosymbioses resulted in the establishment of plastids with two membranes. However, a vast variety of eukaryotes possess plastids with three or more membranes. They derive from the endosymbioses of primary photosynthetic eukaryotes within other eukaryotic cells (Delwiche 1999; Keeling 2013). Such secondary endosymbioses have spread photosynthesis across the eukaryotic tree, either by the endosymbiosis of red or of green algae. Whereas it is almost certain that secondary endosymbioses of green algae occurred twice (in euglenids and chlorarachniophytes), secondary red algal plastids are found in a variety of alveolates, stramenopiles, cryptophytes, and haptophytes, and the number of red algal endosymbioses at the origin of these groups has been matter of intense debate (Baurain et al. 2010; Keeling 2010, 2013; Burki et al. 2012b). Moreover, the existence of tertiary endosymbioses (namely, the symbiosis of a secondary photosynthetic eukaryote within another eukaryotic cell) and of plastid replacements makes the picture of plastid evolution in eukaryotes even more complex. Dinoflagellates, some of which have replaced their ancestral red algal plastids by green algae, diatoms, haptophytes, or cryptophytes, are paradigmatic examples of such complex situations (Keeling 2013).The evolution of plastids has been studied using genes from the plastid genome as well as typical eukaryotic nuclear genes, which allow inferring the phylogenies of both the plastids and their hosts. The use of those markers has led to interesting discoveries, such as the monophyly of the Archaeplastida (Moreira et al. 2000; Rodríguez-Ezpeleta et al. 2005) or the difficulties in reconciling the plastid and host histories in eukaryotes with red algal plastids (Baurain et al. 2010; Burki et al. 2012b). However, a third class of genes can also provide useful complementary information: the genes of plastid origin retrieved within the nuclear genome of the host. In fact, contemporary plastids have small genomes, which is due to the fact that most of the original cyanobacterial symbiont genes were lost or transferred to the host nucleus (by a process called endosymbiotic gene transfer, EGT) during the evolution of plastids (Weeden 1981; Martin et al. 1998). These transfer events are not restricted to plastid endosymbioses—the same phenomenon occurred during the endosymbiosis that gave rise to the mitochondria (Gray et al. 1999; Burger et al. 2003).EGT genes may serve to study the evolutionary history of plastids and, in particular, the presence of cryptic endosymbioses. In fact, species that had a plastid in the past but lost photosynthesis may have conserved genes of plastid origin in their nuclear genomes. This has been shown for a variety of nonphotosynthetic eukaryotes, such as, for example, apicomplexan parasites (Fast et al. 2001; Roos et al. 2002; Williams and Keeling 2003; Huang et al. 2004), perkinsids (Stelter et al. 2007; Matsuzaki et al. 2008; Fernández Robledo et al. 2011) or nonphotosynthetic dinoflagellates (Sanchez-Puerta et al. 2007; Slamovits and Keeling 2008), and green algae (de Koning and Keeling 2004). Although much more controversial, potential EGTs have also been used to propose a photosynthetic ancestry for ciliates (Reyes-Prieto et al. 2008) or that algae with secondary plastids of red algal origin, such as diatoms and chromerids, may have contained green algal endosymbionts in their past (Moustafa et al. 2009; Woehle et al. 2011). Likewise, several dozens of potential EGTs have been detected in algae and plants that appear to have been acquired from Chlamydiae, a group of parasitic bacteria (Huang and Gogarten 2007; Becker et al. 2008; Moustafa et al. 2008), which led to proposing that cryptic chlamydial endosymbionts may have helped to establish the first plastids, in particular, by providing essential functions for plastid activity (Greub and Raoult 2003; Ball et al. 2013; Baum 2013).We revise here some of these cases of cryptic endosymbiosis, with special attention on the difficulties in accurately detecting EGT and the importance of proper phylogenetic analysis and of an adequate taxonomic sampling to achieve that task.     

Cadherin-mediated Intercellular Adhesion and Signaling Cascades Involving Small GTPases

Epithelia form physical barriers that separate the internal milieu of the body from its external environment. The biogenesis of functional epithelia requires the precise coordination of many cellular processes. One of the key events in epithelial biogenesis is the establishment of cadherin-dependent cell–cell contacts, which initiate morphological changes and the formation of other adhesive structures. Cadherin-mediated adhesions generate intracellular signals that control cytoskeletal reorganization, polarity, and vesicle trafficking. Among such signaling pathways, those involving small GTPases play critical roles in epithelial biogenesis. Assembly of E-cadherin activates several small GTPases and, in turn, the activated small GTPases control the effects of E-cadherin-mediated adhesions on epithelial biogenesis. Here, we focus on small GTPase signaling at E-cadherin-mediated epithelial junctions.Cell–cell adhesions are involved in a diverse range of physiological processes, including morphological changes during tissue development, cell scattering, wound healing, and synaptogenesis (Adams and Nelson 1998; Gumbiner 2000; Halbleib and Nelson 2006; Takeichi 1995; Tepass et al. 2000). In epithelial cells, cell–cell adhesions are classified into three kinds of adhesions: adherens junction, tight junction, and desmosome (for more details, see Meng and Takeichi 2009, Furuse 2009, and Delva et al. 2009, respectively). A key event in epithelial polarization and biogenesis is the establishment of cadherin-dependent cell–cell contacts. Cadherins belong to a large family of adhesion molecules that require Ca²⁺ for their homophilic interactions (Adams and Nelson 1998; Blanpain and Fuchs 2009; Gumbiner 2000; Hartsock and Nelson 2008; Takeichi 1995; Tepass et al. 2000). Cadherins form transinteraction on the surface of neighboring cells (for details, see Shapiro and Weis 2009). For the development of strong and rigid adhesions, cadherins are clustered concomitantly with changes in the organization of the actin cytoskeleton (Tsukita et al. 1992). Classical cadherins are required, but not sufficient, to initiate cell–cell contacts, and other adhesion protein complexes subsequently assemble (for details, see Green et al. 2009). These complexes include the tight junction, which controls paracellular permeability, and desmosomes, which support the structural continuum of epithelial cells. A fundamental problem is to understand how these diverse cellular processes are regulated and coordinated. Intracellular signals, generated when cells attach with one another, mediate these complicated processes.Several signaling pathways upstream or downstream of cadherin-mediated cell–cell adhesions have been identified (Perez-Moreno et al. 2003) (see also McCrea et al. 2009). Among these pathways, small GTPases including the Rho and Ras family GTPases play critical roles in epithelial biogenesis and have been studied extensively. Many key morphological and functional changes are induced when these small GTPases act at epithelial junctions, where they mediate an interplay between cell–cell adhesion molecules and fundamental cellular processes including cytoskeletal activity, polarity, and vesicle trafficking. In addition to these small GTPases, Ca²⁺ signaling and phosphorylation of cadherin complexes also play pivotal roles in the formation and maintenance of cadherin-mediated adhesions. Here, we focus on signaling pathways involving the small GTPases in E-cadherin-mediated cell–cell adhesions. Other signaling pathways are described in recent reviews (Braga 2002; Fukata and Kaibuchi 2001; Goldstein and Macara 2007; McLachlan et al. 2007; Tsukita et al. 2008; Yap and Kovacs 2003; see also McCrea et al. 2009).     

Cell Biology of the Endoplasmic Reticulum and the Golgi Apparatus through Proteomics

Enriched endoplasmic reticulum (ER) and Golgi membranes subjected to mass spectrometry have uncovered over a thousand different proteins assigned to the ER and Golgi apparatus of rat liver. This, in turn, led to the uncovering of several hundred proteins of poorly understood function and, through hierarchical clustering, showed that proteins distributed in patterns suggestive of microdomains in cognate organelles. This has led to new insights with respect to their intracellular localization and function. Another outcome has been the critical testing of the cisternal maturation hypothesis showing overwhelming support for a predominant role of COPI vesicles in the transport of resident proteins of the ER and Golgi apparatus (as opposed to biosynthetic cargo). Here we will discuss new insights gained and also highlight new avenues undertaken to further explore the cell biology of the ER and the Golgi apparatus through tandem mass spectrometry.Most biosynthetic proteins destined for the plasma membrane and secretion, as well as resident proteins of endosomes, lysosomes, the endoplasmic reticulum (ER), and the Golgi apparatus originate with cotranslational translocation into the ER. This is followed by carefully controlled folding and quality control. The ER is also a major site for sensing cellular stress and for cholesterol and phospholipid biosynthesis and constitutes a vast continuous endomembrane system that often pervades the entire cytoplasm. The ER commences as the nuclear envelope extends to rough (ribosome-studded) membranes, and ends with tripartite-like structures of tubular smooth (ribosome-free) ER.The ratio of rough and smooth ER is cell type specific. Stem cells have little rough ER and smooth ER (Murphy et al. 1971; Cheng and Leblond 1974), whereas cells highly specialized in protein secretion, such as in the exocrine pancreas, have extensive rough ER with little to no smooth ER (Palade and Siekevitz 1956; Jamieson and Palade 1967). By contrast, liver parenchymal hepatocytes have a near equal abundance of rough and smooth ER with the latter associated with glycogen storage and elimination of exogenous steroids through a multitude of P450-driven oxidation pathways (Bruni and Porter 1965; Loud 1968; Estabrook et al. 1971; Blouin et al. 1977; Reed and Backes 2012). At the other extreme, Leydig cells of the testis are specialized in cholesterol metabolism to form testosterone and are enriched with smooth ER (Mori and Christensen 1980). Emanating from the ER of all cell types in mammals are discrete export sites that are distributed throughout the cell, sometimes numbering in the hundreds. ER export is controlled by the COPII coat machinery, which has been characterized through proteomics analyses in yeast (Otte et al. 2001) and is coupled to microtubule- and dynein/dynactin-dependent transport directed toward the central juxta-nuclear Golgi apparatus (COPII vesicles are discussed in Lord et al. [2013]). This occurs in vesicular/tubular clusters (VTCs) that seemingly undergo a maturation/distillation process (Saraste and Kuismanen 1984) such that when arriving at the cis-face Golgi stacks, biosynthetic cargo appears more concentrated. This is achieved, at least in part, through a continuous removal of ER export machinery components (e.g., p58, p24, and SNARE proteins) that are then returned to the ER via vesicular transport intermediates controlled by the COPI coat machinery. This apparent “distillation” process continues throughout the secretory pathway from as far as the trans-part of the Golgi apparatus (Miesenbock and Rothman 1995). Whether or not such distillation occurs through intra-Golgi cisternal transport or through direct delivery to the ER is part of ongoing investigation, including the extent of recycling (see below).The Golgi apparatus is morphologically distinct from the ER because of its juxtanuclear position, and at the ultrastructural level, appears as a ribbon-like structure of laterally interconnected stacks of flattened cisternae, each having a network of extensive fenestrated membranes associated both at their cis- and trans-face. Biosynthetic cargo here, undergo extensive posttranslational modifications including the maturation of N-linked oligosaccharides, and the addition of O-linked oligosaccharides. At the trans-face, cargo is sorted and packaged for transport either to the plasma membrane or to the endosomal endomembrane system. Some specialized cargo is also packaged and concentrated into dedicated membrane structures for regulated secretion.The functional and morphological demarcation between the ER and the Golgi apparatus is usually assumed to be complete, giving rise to the notion of two independent organelles—each with their own distinct functions. Because of the extensive recycling that takes place, however, the two organelles appear functionally intertwined. Indeed, inhibition of the Golgi-located ARF1 guanine nucleotide exchange factor GBF1 results in a rapid collapse of the Golgi apparatus into the ER (Misumi et al. 1986; Oda et al. 1987; Fujiwara et al. 1988; Lippincott-Schwartz et al. 1989; Claude et al. 1999). As ARF1^GDP to ARF1^GTP conversion is obligatory to COPI recruitment and vesicle formation and subsequent recycling, the effect of BFA seems at first counter intuitive, but can be explained through the additional role of COPI as a protective coat on cisternal membranes, i.e., BFA-induced dissociation of COPI coat from Golgi membranes enables uncontrolled formation of Golgi-derived tubules that within minutes fuse with the ER, collapsing most of the Golgi into the ER. Strikingly, removal of BFA equally rapidly leads to the reformation of individual functional Golgi stacks positioned at the various ER exit sites and after microtubule-dependent transport, coalesce into the central and juxtanuclear Golgi apparatus. Thus, the two organelles are temporally as well as spatially linked to enable productive communication and trafficking for the process of protein secretion, as well as lipid biosynthesis. This link gives rise to some unexpected functions (see below).