Widely Conserved Signaling Pathways in the Establishment of Cell Polarity

How are the asymmetric distributions of proteins, lipids, and RNAs established and maintained in various cell types? Studies from diverse organisms show that Par proteins, GTPases, kinases, and phosphoinositides participate in conserved signaling pathways to establish and maintain cell polarity.The asymmetric distribution of proteins, lipids, and RNAs is necessary for cell fate determination, differentiation, and specialized cell functions that underlie morphogenesis (St Johnston 2005; Gonczy 2008; Knoblich 2008; Macara and Mili 2008; Martin-Belmonte and Mostov 2008). A fundamental question is how this asymmetric distribution is established and maintained in different types of cells and tissues. The formation of a specialized apical surface on an epithelial cell seems quite different from the specification of axons versus dendrites in a neuron, or the asymmetric division of a nematode zygote. Yet, remarkably, a conserved molecular toolbox is used throughout the metazoa to establish and maintain cell polarity in these and many other contexts. This toolbox consists of proteins that are components of signal transduction pathways (Goldstein and Macara 2007; Assemat et al. 2008; Yamanaka and Ohno 2008). However, our understanding of these pathways, and their intersection with other signaling networks, remains incomplete. Moreover, the regulation and cross talk between the polarity proteins and other signaling components varies from one context to another, which complicates the task of dissecting polarity protein function. Nonetheless, rapid progress is being made in our understanding of polarity signaling, which is outlined in this article, with an emphasis on the Par proteins, because these proteins play major roles integrating diverse signals that regulate cell polarity (Fig. 1) (see Munro and Bowerman 2009; Prehoda 2009; Nelson 2009).Open in a separate windowFigure 1.An overview of Par complex signaling, showing inputs (bottom) and outputs (top) with cellular functions that are targeted by these pathways (italics).     

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).     

Diversity of Neural Precursors in the Adult Mammalian Brain

Aided by advances in technology, recent studies of neural precursor identity and regulation have revealed various cell types as contributors to ongoing cell genesis in the adult mammalian brain. Here, we use stem-cell biology as a framework to highlight the diversity of adult neural precursor populations and emphasize their hierarchy, organization, and plasticity under physiological and pathological conditions.The adult mammalian brain displays remarkable structural plasticity by generating and incorporating new neural cell types into an already formed brain (Kempermann and Gage 1999). Largely restricted within the subventricular zone (SVZ) along the lateral ventricle and the subgranular zone (SGZ) in the dentate gyrus (DG), neural genesis is thought to arise from neural stem cells (NSCs) (Ming and Song 2011). Stem cells are defined by hallmark functions: capacity to self-renew, maintenance of an immature state over a long duration, and ability to generate specialized cell types (Fig. 1). These features distinguish stem cells from committed progenitor cells that more readily differentiate into specialized cell types (Fig. 1). Stem and progenitor cells (collectively called precursors) are additionally characterized by their lineage capacity. For example, multipotential neural precursors generate neurons and glia, whereas unipotential cells produce only one cell type, such as neurons (Gage 2000; Ma et al. 2009). The classical NSC definition is based on cell culture experiments in which a single cell can self-renew and generate neurons, astrocytes, and oligodendrocytes (Gage 2000; Ma et al. 2009). Yet, reprogramming studies have raised the question of whether cultured lineage-restricted neural progenitors acquire additional potential not evident in vivo (Palmer et al. 1999; Kondo and Raff 2000; Gabay et al. 2003). As a result, various lineage models have been proposed to explain cell generation in the adult brain (Fig. 1) (Ming and Song 2011). In one model, bona fide adult stem cells generate multiple lineages at the individual cell level. In another, cell genesis represents a collective property from a mixed population of unipotent progenitors. Importantly, these models are not mutually exclusive as evidence for the coexistence of multiple precursors has been observed in several adult somatic tissues, in which one population preferentially maintains homeostasis and another serves as a cellular reserve (Li and Clevers 2010; Mascre et al. 2012). Recent technical advances, including single-cell lineage tracing (Kretzschmar and Watt 2012), have made it possible to dissect basic cellular and behavioral processes of neural precursors in vivo (Fig. 4) (Bonaguidi et al. 2012). In this work, we review our current knowledge of precursor cell identity, hierarchical organization, and regulation to examine the diverse origins of cell genesis in the adult mammalian brain.Open in a separate windowFigure 1.Models of generating cell diversity in the adult tissues. (A,B) Definitions of stem and progenitor cells. In A, quiescent stem cells (S_q) become active stem cells (S_a) that proliferate to generate different types of specialized cells (C₁, C₂, C₃) and new stem cells (S). The active stem cell can return to quiescence and remain quiescent over long periods of time. In B, lineage-restricted progenitor cells lacking self-renewal capacity (P₁, P₂, P₃) each give rise to distinct populations of specialized cells (C₁, C₂, C₃). (C) Generation of specialized cells in a tissue could be explained by three models. (1) The stem-cell model, in which multipotent stem cells give rise to all the specialized cells in the tissue. (2) The progenitor cell model, in which diverse, lineage-restricted progenitor cells give rise to different cell types in the tissue. (3) A hybrid model, in which a mixture of stem cells and lineage-restricted progenitor cells generate specialized cells of the adult tissue.

Table 1.

Comparison of different methods used to study the generation of new cells in the adult mammalian nervous system

Poles Apart: Prokaryotic Polar Organelles and Their Spatial Regulation

While polar organelles hold the key to understanding the fundamentals of cell polarity and cell biological principles in general, they have served in the past merely for taxonomical purposes. Here, we highlight recent efforts in unraveling the molecular basis of polar organelle positioning in bacterial cells. Specifically, we detail the role of members of the Ras-like GTPase superfamily and coiled-coil-rich scaffolding proteins in modulating bacterial cell polarity and in recruiting effector proteins to polar sites. Such roles are well established for eukaryotic cells, but not for bacterial cells that are generally considered diffusion-limited. Studies on spatial regulation of protein positioning in bacterial cells, though still in their infancy, will undoubtedly experience a surge of interest, as comprehensive localization screens have yielded an extensive list of (polarly) localized proteins, potentially reflecting subcellular sites of functional specialization predicted for organelles.Since the first electron micrographs that revealed flagella at the cell poles of bacteria, we have known that bacterial cells are polarized and that they are able to decode the underlying positional information to confine the assembly of an extracellular organelle to a polar cellular site (Fig. 1). Foraging into this unknown territory has been challenging, but recent efforts that exploit the power of bacterial genetics along with modern imaging methods to visualize proteins in the minute bacterial cells has yielded several enticing entry points to dissect polarity-based mechanisms and explore potentially contributing subdiffusive characteristics (Golding and Cox 2006).Open in a separate windowFigure 1.Transmission electron micrograph (taken by Jeff Skerker) of a Caulobacter crescentus swarmer cell showing the polar pili (empty arrowheads), the polar flagellum with the flagellar filament (filled arrowheads), and the hook (white arrow) (see Fig. 2A).While polar organelles are a visual manifestation of polarity, it is important to point out that polarity can also be inherent to cells, at least in molecular terms, even in the absence of discernible polar structures. In other words, molecular anatomy can reveal that a bacterial cell, such as an Escherichia coli cell, features specialized protein complexes at or near the poles, despite a perfectly symmetrical morphology (Maddock and Shapiro 1993; Lindner et al. 2008). Such systemic polarization in bacteria, likely stemming from the distinctive division history of each pole, has the potential to be widespread and to be exploited for positioning of polar organelles and protein complexes. As excellent reviews have been published detailing the interplay between cell polarity and protein localization (Dworkin 2009; Shapiro et al. 2009; Kaiser et al. 2010; Rudner and Losick 2010), here we focus on recent progress in understanding the function and localization of spatial regulators of polar organelles. Considering that the ever-growing list of polar protein complexes emerging from systematic and comprehensive localization studies (Kitagawa et al. 2005; Russell and Keiler 2008; Werner et al. 2009; Hughes et al. 2010) is suggestive of multiple polarly confined (organelle-like) functions, understanding their spatial regulation is also of critical relevance in the realm of medical bacteriology, as many virulence determinants also underlie polarity (Goldberg et al. 1993; Scott et al. 2001; Judd et al. 2005; Jain et al. 2006; Jaumouille et al. 2008; Carlsson et al. 2009). Below, we highlight a few prominent examples of overtly polar organelles and the proteins known to date that regulate their polar positioning.     

Toll-Like Receptor Signaling

Toll-like receptors sense pathogen-associated molecular patterns (e.g., lipopolysaccharides) and trigger gene-expression changes that ultimately eradicate the invading microbes.Toll-like receptors (TLRs) are protective immune sentries that sense pathogen-associated molecular patterns (PAMPs) such as unmethylated double-stranded DNA (CpG), single-stranded RNA (ssRNA), lipoproteins, lipopolysaccharide (LPS), and flagellin. In innate immune myeloid cells, TLRs induce the secretion of inflammatory cytokines (Newton and Dixit 2012), thereby engaging lymphocytes to mount an adaptive, antigen-specific immune response (see Fig. 1) that ultimately eradicates the invading microbes (Kawai and Akira 2010).Open in a separate windowFigure 1.TLR signaling (simplified view).Identification of TLR innate immune function began with the discovery that Drosophila mutants in the Toll gene are highly susceptible to fungal infection (Lemaitre et al. 1996). This was soon followed by identification of a human Toll homolog, now known as TLR4 (Medzhitov et al. 1997). To date, 10 TLR family members have been identified in humans, and at least 13 are present in mice. All TLRs consist of an amino-terminal domain, characterized by multiple leucine-rich repeats, and a carboxy-terminal TIR domain that interacts with TIR-containing adaptors. Nucleic acid–sensing TLRs (TLR3, TLR7, TLR8, and TLR9) are localized within endosomal compartments, whereas the other TLRs reside at the plasma membrane (Blasius and Beutler 2010; McGettrick and O’Neill 2010). Trafficking of most TLRs from the endoplasmic reticulum (ER) to either the plasma membrane or endolysosomes is orchestrated by ER-resident proteins such as UNC93B (for TLR3, TLR7, TLR8, and TLR9) and PRAT4A (for TLR1, TLR2, TLR4, TLR7, and TLR9) (Blasius and Beutler 2010). Once in the endolysosomes, TLR3, TLR7, and TLR9 are subject to stepwise proteolytic cleavage, which is required for ligand binding and signaling (Barton and Kagan 2009). For some TLRs, ligand binding is facilitated by coreceptors, including CD14 and MD2.Following ligand engagement, the cytoplasmic TIR domains of the TLRs recruit the signaling adaptors MyD88, TIRAP, TRAM, and/or TRIF (see Fig. 2). Depending on the nature of the adaptor that is used, various kinases (IRAK4, IRAK1, IRAK2, TBK1, and IKKε) and ubiquitin ligases (TRAF6 and pellino 1) are recruited and activated, culminating in the engagement of the NF-κB, type I interferon, p38 MAP kinase (MAPK), and JNK MAPK pathways (Kawai and Akira 2010; Morrison 2012). TRAF6 is modified by K63-linked autoubiquitylation, which enables the recruitment of IκB kinase (IKK) through a ubiquitin-binding domain of the IKKγ (also known as NEMO) subunit. In addition, a ubiquitin-binding domain of TAB2 recognizes ubiquitylated TRAF6, causing activation of the associated TAK1 kinase, which then phosphorylates the IKKβ subunit. Pellino 1 can modify IRAK1 with K63-linked ubiquitin, allowing IRAK1 to recruit IKK directly. TLR4 signaling via the TRIF adaptor protein leads to K63-linked polyubiquitylation of TRAF3, thereby promoting the type I interferon response via interferon regulatory factor (IRFs) (Hacker et al. 2011). Alternatively, TLR4 signaling via MyD88 leads to the activation of TRAF6, which modifies cIAP1 or cIAP2 with K63-linked polyubiquitin (Hacker et al. 2011). The cIAPs are thereby activated to modify TRAF3 with K48-linked polyubiquitin, causing its proteasomal degradation. This allows a TRAF6–TAK1 complex to activate the p38 MAPK pathway and promote inflammatory cytokine production (Hacker et al. 2011). TLR signaling is turned off by various negative regulators: IRAK-M and MyD88 short (MyD88s), which antagonize IRAK1 activation; FADD, which antagonizes MyD88 or IRAKs; SHP1 and SHP2, which dephosphorylate IRAK1 and TBK1, respectively; and A20, which deubiquitylates TRAF6 and IKK (Flannery and Bowie 2010; Kawai and Akira 2010).Open in a separate windowFigure 2.TLR signaling. (Adapted with kind permission of Cell Signaling Technology [http://www.cellsignal.com].)Deregulation of the TLR signaling cascade causes several human diseases. Patients with inherited deficiencies of MyD88, IRAK4, UNC93B1, or TLR3 are susceptible to recurrent bacterial or viral infections (Casanova et al. 2011). Chronic TLR7 and/or TLR9 activation in autoreactive B cells, in contrast, underlies systemic autoimmune diseases (Green and Marshak-Rothstein 2011). Furthermore, oncogenic activating mutations of MyD88 occur frequently in the activated B-cell-like subtype of diffuse large B-cell lymphoma and in other B-cell malignancies (Ngo et al. 2011). Inhibitors of various TLRs or their associated kinases are currently being developed for autoimmune or inflammatory diseases and also hold promise for the treatment of B-cell malignancies with oncogenic MyD88 mutations. Many TLR7 and TLR9 agonists are currently in clinical trials as adjuvants to boost host antitumor responses in cancer patients (Hennessy et al. 2010).     

Auxin and Monocot Development

Monocots are known to respond differently to auxinic herbicides; hence, certain herbicides kill broadleaf (i.e., dicot) weeds while leaving lawns (i.e., monocot grasses) intact. In addition, the characters that distinguish monocots from dicots involve structures whose development is controlled by auxin. However, the molecular mechanisms controlling auxin biosynthesis, homeostasis, transport, and signal transduction appear, so far, to be conserved between monocots and dicots, although there are differences in gene copy number and expression leading to diversification in function. This article provides an update on the conservation and diversification of the roles of genes controlling auxin biosynthesis, transport, and signal transduction in root, shoot, and reproductive development in rice and maize.Auxinic herbicides have been used for decades to control dicot weeds in domestic lawns (Fig. 1A), commercial golf courses, and acres of corn, wheat, and barley, yet it is not understand how auxinic herbicides selectively kill dicots and spare monocots (Grossmann 2000; Kelley and Reichers 2007). Monocots, in particular grasses, must perceive or respond differently to exogenous synthetic auxin than dicots. It has been proposed that this selectivity is because of either limited translocation or rapid degradation of exogenous auxin (Gauvrit and Gaillardon 1991; Monaco et al. 2002), altered vascular anatomy (Monaco et al. 2002), or altered perception of auxin in monocots (Kelley and Reichers 2007). To explain these differences, there is a need to further understand the molecular basis of auxin metabolism, transport, and signaling in monocots.Open in a separate windowFigure 1.Differences between monocots and dicots. (A) A dicot weed in a lawn of grasses. Note the difference in morphology of the leaves. (B) Germinating dicot (bean) seedling. Dicots have two cotyledons (cot). Reticulate venation is apparent in the leaves. The stem below the cotyledons is called the hypocotyl (hyp). (C) Germinating monocot (maize) seedling. Monocots have a single cotyledon called the coleoptile (col) in grasses. Parallel venation is apparent in the leaves. The stem below the coleoptile is called the mesocotyl (mes).Auxin, as we have seen in previous articles, plays a major role in vegetative, reproductive, and root development in the model dicot, Arabidopsis. However, monocots have a very different anatomy from dicots (Raven et al. 2005). Many of the characters that distinguish monocots and dicots involve structures whose development is controlled by auxin: (1) As the name implies, monocots have single cotyledons, whereas dicots have two cotyledons (Fig. 1B,C). Auxin transport during embryogenesis may play a role in this difference as cotyledon number defects are often seen in auxin transport mutants (reviewed in Chandler 2008). (2) The vasculature in leaves of dicots is reticulate, whereas the vasculature in monocots is parallel (Fig. 1). Auxin functions in vascular development because many mutants defective in auxin transport, biosynthesis, or signaling have vasculature defects (Scarpella and Meijer 2004). (3) Dicots often produce a primary tap root that produces lateral roots, whereas, in monocots, especially grasses, shoot-borne adventitious roots are the most prominent component of the root system leading to the characteristic fibrous root system (Fig. 2). Auxin induces lateral-root formation in dicots and adventitious root formation in grasses (Hochholdinger and Zimmermann 2008).Open in a separate windowFigure 2.The root system in monocots. (A) Maize seedling showing the primary root (1yR), which has many lateral roots (LR). The seminal roots (SR) are a type of adventitious root produced during embryonic development. Crown roots (CR) are produced from stem tissue. (B) The base of a maize plant showing prop roots (PR), which are adventitious roots produced from basal nodes of the stem later in development.It is not yet clear if auxin controls the differences in morphology seen in dicots versus monocots. However, both conservation and diversification of mechanisms of auxin biosynthesis, homeostasis, transport, and signal transduction have been discovered so far. This article highlights the similarities and the differences in the role of auxin in monocots compared with dicots. First, the genes in each of the pathways are introduced (Part I, Table I) and then the function of these genes in development is discussed with examples from the monocot grasses, maize, and rice (Part II).     

Central Role of RET in Thyroid Cancer

RET (rearranged during transfection) is a receptor tyrosine kinase involved in the development of neural crest derived cell lineages, kidney, and male germ cells. Different human cancers, including papillary and medullary thyroid carcinomas, lung adenocarcinomas, and myeloproliferative disorders display gain-of-function mutations in RET. Accordingly, RET protein has become a promising molecular target for cancer treatment.The human RET (rearranged during transfection) gene maps on 10q11.2 and is composed of 21 exons spanning a region of 55,000 bp. It encodes a single-pass trans-membrane protein, RET, that belongs to the receptor tyrosine kinase (RTK) family (Pasini et al. 1995). The RET extracellular segment contains four cadherin-like domains, followed by a domain containing cysteine residues involved in the formation of intramolecular disulfide bonds (Fig. 1A) (Anders et al. 2001; Airaksinen and Saarma 2002). RET protein is highly glycosylated and N-glycosylation is necessary for its transport to the cell surface. Only the fully mature glycosylated 170 kDa RET protein isoform is exposed to the extracellular compartment, whereas the mannose-rich 150 kDa isoform is confined to the Golgi (Takahashi et al. 1993; Carlomagno et al. 1996). The transmembrane segment is composed of 22 amino acids, among which S649 and S653 mediate self-association and dimerization of RET, possibly via formation of inter-molecular hydrogen bonding (Kjaer et al. 2006). The intracellular portion of RET contains the tyrosine kinase domain split into two subdomains by the insertion of 27 amino acids. The RET COOH-terminal tail varies in length as a result of alternative splicing of the 3′ end (carboxy terminal with respect to glycine 1063), generating three different isoforms that contain 9 (RET9), 43 (RET43), or 51 (RET51) amino acids (Myers et al. 1995). RET9 and RET51 are the most abundant isoforms, and they activate similar signaling pathways through interaction with diverse protein complexes, and may exert a differential role in development (Fig. 1A) (de Graaff et al. 2001).Open in a separate windowFigure 1.Illustration of the mechanisms of activation of wild-type (wt) RET and RET-derived oncoproteins. (A) Wild-type RET activation is mediated by ligand (GFL)-induced dimerization; ligand binding to RET is not direct and mediated by GFR-α coreceptors (not shown); major RET autophosphorylation sites and downstream signaling pathways are indicated. RET extracellular cadherin-like domains are represented in red. The split intracellular RET tyrosine kinase domain, as well as the three alternative carboxy-terminal RET tails, are also depicted. (B) RET/PTC activation is mediated by coiled-coil-induced dimerization (left); activation of RET cysteine mutants associated with MEN2A or FMTC is mediated by disulfide bonds-mediated dimerization (right).RET shows several autophosphorylation sites (Fig. 1A) (Liu et al. 1996; Kawamoto et al. 2004). RET tyrosine 1062 (Y1062) functions as a multidocking site for signaling molecules containing a phosphotyrosine-binding (PTB) domain (Asai et al. 1996). Phospho-Y1062 binding proteins include SHC, N-SHC (RAI), FRS2, IRS1/2, DOK1, and DOK4/5 that, in turn, contribute to the activation of RAS-MAPK (mitogen-activated protein kinases) and PI3K (phosphatidyl inositol 3 kinase)-AKT pathways. Y1096, specific to the RET51 splicing variant, couples to the PI3K-AKT and RAS-MAPK pathways, as well. These signaling cascades mediate RET-dependent cell survival, proliferation, and motility (Alberti et al. 1998; Murakami et al. 1999; Segouffin-Cariou and Billaud 2000; Melillo et al. 2001a,b; Schuetz et al. 2004). Y905 is located in the activation loop of the RET kinase and its phosphorylation is associated with RET kinase activation (Knowles et al. 2006). Finally, Y981 and Y1015 have been shown to be coupled to important signaling molecules such as SRC and PLC-γ, respectively (Borrello et al. 1996; Encinas et al. 2004).RET is the receptor for a group of neurotrophic growth factors that belong to the glial cell line-derived neurotrophic factor (GDNF) family (GFLs, GDNF family ligands), namely, GDNF, Neurturin (NRT), Artemin (ART), and Persephin (PSF) (Airaksinen and Saarma 2002). GFLs mediate RET protein dimerization and activation (Fig. 1A). GFLs are presented to RET by GPI (glycosylphosphatidylinositol)-anchored coreceptors, called GFR-α (GDNF family receptor α 1-4). Differential tissue expression dictates the specificity of action displayed by alternative GLF-GFR-α pairs during development and adult life (Baloh et al. 2000; Airaksinen and Saarma 2002).Together with other membrane (DCC and p75NTR) or nuclear (androgen receptor, AR) receptors, RET belongs to the family of so-called “dependence” receptors (Mehlen and Bredesen 2011). In the absence of ligand, RET exerts a proapoptotic activity, that is blocked on ligand stimulation (Bordeaux et al. 2000). Such pro-apoptotic activity is RET kinase-independent and mediated by cleavage of RET cytosolic portion by caspase-3, which, in turn, releases a carboxy-terminal RET peptide that is able to induce cell death (Bordeaux et al. 2000). It is feasible that such activity is important for RET developmental function, because it may control migration of RET-expressing cells by limiting survival of cells that move beyond ligand availability (Bordeaux et al. 2000; Cañibano et al. 2007). Whether modulation of this function is also important for RET-associated diseases is still unknown. However, it is interesting to note that a cancer-associated RET mutant (RET-C634R, see below) does not exert cleavage-dependent proapoptotic effects, whereas RET mutants associated with defective development (Hirschsprung disease, see below) exert strong proapoptotic activity that is refractory to modulation by ligand (Bordeaux et al. 2000).RET is expressed in enteric ganglia, adrenal medulla chromaffin cells, thyroid C cells, sensory and autonomic ganglia of the peripheral nervous system, a subset of central nervous system nuclei, developing kidney and testis germ cells (Manié et al. 2001; de Graaff et al. 2001). RET null mice display impaired development of superior cervical ganglia and enteric nervous system, kidney agenesia, reduction of thyroid C cells, and impaired spermatogenesis (Manié et al. 2001). Accordingly, individuals with germline loss-of-function mutations of RET are affected by intestinal aganglionosis causing congenital megacolon (Hirschsprung disease) (Brooks et al. 2005). RET loss-of-function mutations have also been identified in congenital anomalies of kidney and urinary tract (CAKUT), either isolated or in combination with Hirschsprung disease (Jain 2009).Several genetic alterations convert RET into a dominantly transforming oncogene. This review will describe RET-derived oncogenes that are associated with different types of human neoplasia (Fig. 1B).     

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.     

Membrane Domains Based on Ankyrin and Spectrin Associated with Cell–Cell Interactions

Nodes of Ranvier and axon initial segments of myelinated nerves, sites of cell–cell contact in early embryos and epithelial cells, and neuromuscular junctions of skeletal muscle all perform physiological functions that depend on clustering of functionally related but structurally diverse ion transporters and cell adhesion molecules within microdomains of the plasma membrane. These specialized cell surface domains appeared at different times in metazoan evolution, involve a variety of cell types, and are populated by distinct membrane-spanning proteins. Nevertheless, recent work has shown that these domains all share on their cytoplasmic surfaces a membrane skeleton comprised of members of the ankyrin and spectrin families. This review will summarize basic features of ankyrins and spectrins, and will discuss emerging evidence that these proteins are key players in a conserved mechanism responsible for assembly and maintenance of physiologically important domains on the surfaces of diverse cells.Spectrins are flexible rods 0.2 microns in length with actin-binding sites at each end (Shotton et al. 1979; Bennett et al. 1982) (Fig. 1A). Spectrins are assembled from α and β subunits, each comprised primarily of multiple copies of a 106-amino acid repeat (Speicher and Marchesi 1984). In addition to the canonical 106-residue repeat, β spectrins also have a carboxy-terminal pleckstrin homology domain (Zhang et al. 1995; Macias et al. 1994) and tandem amino-terminal calponin homology domains (Bañuelos et al. 1998), whereas α spectrins contain an Src homology domain 3 (SH3) site (Musacchio et al. 1992), a calmodulin-binding site (Simonovic et al. 2006), and EF hands (Travé et al. 1995) (Fig. 1A). Spectrin α and β subunits are assembled antiparallel and side-to-side into heterodimers, which in turn are associated head-to-head to form tetramers (Clarke 1971; Shotton et al. 1979; Davis and Bennett 1983) (Fig. 1A). In human erythrocytes, in which spectrin was first characterized (Marchesi and Steers 1968; Clarke 1971), actin oligomers containing 10–14 monomers are each linked to five to six spectrin tetramers by accessory proteins to form a geodesic domelike structure that has been resolved by electron microscopy (Byers and Branton 1985). The principal proteins at the spectrin–actin junction are protein 4.1, adducin, tropomyosin, tropomodulin, and dematin (Bennett and Baines 2001) (Open in a separate windowFigure 1.Domain structure and variants of spectrin and ankyrin proteins. (A) Molecular domains of spectrins: Two α spectrins and five β spectrins are shown. Spectrins are comprised of modular units called spectrin repeats (yellow). Other domains such as the ankyrin binding domain (purple), Src-homology domain 3 (SH3, blue), EF-hand domain (red), and calmodulin-binding domain (green) promote interactions with binding targets important for spectrin function. The pleckstrin homology domain (black) promotes association with the plasma membrane and the actin binding domain (grey) tethers the spectrin-based membrane skeleton to the underlying actin cytoskeleton. (B) The spectrin tetramer, the fundamental unit of the spectrin-based membrane skeleton. The spectrin repeat domains of α and β spectrin associate end-to-end to form heterodimers. Heterodimers associate laterally in an antiparallel fashion to form tetramers. The tetramers can then associate end-to-end to form extended macromolecules that link into a geodesic dome shape directly underneath the plasma membrane. (C) Molecular domains present in canonical ankyrins. The membrane binding domain of ankyrin isoforms (orange) is comprised of 24 ANK repeats. The spectrin binding domain (green-blue) allows ankyrins to coordinate integral membrane proteins to the membrane skeleton. The death domain (pink) is the most highly conserved domain. The regulatory domain (brown) is the most variable region of ankyrins. The regulatory domain interacts intramolecularly with the membrane binding domain to modulate ankyrin’s affinity for other binding partners. All ankyrins and spectrins are subject to alternative splicing, which further increases their functional diversity.

Table 1.

Binding partners of spectrin and ankyrins

mGluR1/TRPC3-mediated Synaptic Transmission and Calcium Signaling in Mammalian Central Neurons

Metabotropic glutamate receptors type 1 (mGluR1s) are required for a normal function of the mammalian brain. They are particularly important for synaptic signaling and plasticity in the cerebellum. Unlike ionotropic glutamate receptors that mediate rapid synaptic transmission, mGluR1s produce in cerebellar Purkinje cells a complex postsynaptic response consisting of two distinct signal components, namely a local dendritic calcium signal and a slow excitatory postsynaptic potential. The basic mechanisms underlying these synaptic responses were clarified in recent years. First, the work of several groups established that the dendritic calcium signal results from IP₃ receptor-mediated calcium release from internal stores. Second, it was recently found that mGluR1-mediated slow excitatory postsynaptic potentials are mediated by the transient receptor potential channel TRPC3. This surprising finding established TRPC3 as a novel postsynaptic channel for glutamatergic synaptic transmission.Glutamate is the predominant neurotransmitter used by excitatory synapses in the mammalian brain (Hayashi 1952; Curtis et al. 1959). At postsynaptic sites, glutamate binds to two different classes of receptors, namely the ionotropic glutamate receptors (iGluRs) and the metabotropic glutamate receptors (mGluRs) (Sladeczek et al. 1985; Nicoletti et al. 1986; Sugiyama et al. 1987). The iGluRs represent ligand-gated nonselective cation channels that underlie excitatory postsynaptic currents (EPSCs). Based on their subunit composition, gating, and permeability properties, they are subdivided into three groups named after specific agonists: AMPA- (α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid), NMDA receptors (N-methyl D-aspartate receptors) and kainate receptors (Alexander et al. 2009). The other class of glutamate receptors, the mGluRs, consists of receptors that are coupled to G proteins and act through distinct downstream signaling cascades. They are structurally different from iGluRs and characterized by the presence of seven transmembrane domains (Houamed et al. 1991; Masu et al. 1991). The mGluRs exist as homodimers that do not by themselves form an ion-permeable pore in the membrane (Ozawa et al. 1998). To date, eight different genes (and more splice variants) encoding mGluRs have been identified and form the mGluR1 through mGluR8 subtypes (Alexander et al. 2009). Based on the amino acid sequence homology, downstream signal transduction pathways, and pharmacological properties, each of the subtypes was assigned to one of three groups. Group I receptors consist of mGluR1 and mGluR5 that positively couple to the phospholipase C (PLC). The receptors mGluR2 and mGluR3 constitute group II, whereas the remaining mGluRs, namely mGluR4, mGluR6, mGluR7, and mGluR8, belong to group III. Both groups II and III inhibit the adenylyl cyclase and thereby reduce the concentration of cAMP in the cytosol.Of all different subtypes, mGluR1 is the most abundantly expressed mGluR in the mammalian central nervous system. In the brain, mGluR1 is highly expressed in the olfactory bulb, dentate gyrus, and cerebellum (Lein et al. 2007). The highest expression level of mGluR1 in the brain is found in Purkinje cells, the principal neurons of the cerebellar cortex (Shigemoto et al. 1992; Lein et al. 2007). Together with the AMPA receptors, mGluR1s are part of the excitatory synapses formed between parallel fibers and Purkinje cells (Fig. 1A). Each Purkinje cell is innervated by 100,000–200,000 parallel fibers (Ito 2006) that are axons of the cerebellar granule cells, the most abundant type of neuron in the brain. A second type of excitatory input to Purkinje cells is represented by the climbing fibers that originate in the inferior olive in the brain stem (Ito 2006). The two excitatory synaptic inputs to Purkinje cells are important determinants for the main functions of the cerebellum, including the real-time control of movement precision, error-correction, and control of posture as well as the procedural learning of complex movement sequences and conditioned responses.Open in a separate windowFigure 1.Parallel fiber-evoked mGluR1-dependent signals. (A) Diagram showing the parallel fiber synaptic input to Purkinje cell dendrites. (B) Microelectrode recording of glutamatergic postsynaptic potentials from a Purkinje cell in an acute slice of adult rat cerebellum. Short trains of stimuli to the parallel fibers (5–6 at 50 Hz) caused summation of the early AMPA receptor-dependent EPSPs (leading to spike firing) and a slow, delayed, depolarizing potential (slow EPSP), which was reversibly inhibited by antagonist of mGluRs (+)-MCPG (1mM). (C) Confocal image of a patch-clamped Purkinje cell in a cerebellar slice of an adult mouse. The patch-clamp pipette and the glass capillary used for electrical stimulation of parallel fibers are depicted schematically. The site of stimulation is shown at higher magnification in D. (D) Left: Parallel fiber-evoked (five pulses at 200 Hz, in 10 mM CNQX) synaptic responses consisting of a dendritic mGluR1-dependent Ca²⁺ transient (ΔF/F, top) and an early rapid and a slow excitatory postsynaptic current (EPSC, bottom). Block of the mGluR1-dependent components by the group I-specific mGluR-antagonist CPCCOEt (200 µM) is shown as indicated. Right: Pseudocolor image of the synaptic Ca²⁺ signal. (B, Reprinted with modifications, with permission, from Batchelor and Gaithwaite 1997 [Nature Publishing Group].)It is expected that mGluR1 is involved in many of these cerebellar functions. This view is supported by the observation that mGluR1-deficient knockout mice show severe impairments in motor coordination. In particular, the gait of these mice is strongly affected as well as their ability for motor learning and general coordination (Aiba et al. 1994). The phenotype of the general mGluR1-knockout mice is rescued by the insertion of the gene encoding mGluR1 exclusively into cerebellar Purkinje cells (Ichise et al. 2000) and blockade of mGluR1 expression only in Purkinje cells of adult mice leads to impaired motor coordination (Nakao et al. 2007). These findings established mGluR1 in Purkinje cell as synaptic receptors that are indispensable for a normal cerebellar function.Synaptic transmission involving mGluR1s is found at both parallel fiber-Purkinje cell synapses (Batchelor and Garthwaite 1993; Batchelor et al. 1994) as well as at climbing fiber-Purkinje cell synapses (Dzubay and Otis 2002). Most of our knowledge on the mGluR1 was gained from the analysis of the parallel fiber synapses. The parallel fiber synapse is quite unique in the central nervous system regarding its endowment with neurotransmitter receptors. In contrast to most other glutamatergic synapses in the mammalian brain, it lacks functional NMDA receptors (Shin and Linden 2005). The entire synaptic transmission at these synapses relies on AMPA receptors and on mGluR1 (Takechi et al. 1998). Although AMPA receptors are effectively activated even with single shock stimuli (Konnerth et al. 1990; Llano et al. 1991b), activation of mGluRs requires repetitive stimulation (Batchelor and Garthwaite 1993; Batchelor et al. 1994; Batchelor and Garthwaite 1997; Takechi et al. 1998). A possible explanation for the need of repetitive stimulation may relate to the observation that mGluR1s are found mostly at the periphery of the subsynaptic region (Nusser et al. 1994). At these sites outside the synaptic cleft, glutamate levels that are sufficiently high for receptor activation may be reached only with repetitive stimulation.At parallel fiber-Purkinje cell synapses, repetitive stimulation produces an initial AMPA receptor postsynaptic signal component, followed by a more prolonged mGluR1 component (Fig. 1). Figure 1B shows a current clamp recording of this response consisting of an early burst of action potentials, followed by a prolonged depolarization known as a “slow excitatory postsynaptic potential” (slow EPSP) (Batchelor and Garthwaite 1993; Batchelor et al. 1994; Batchelor and Garthwaite 1997). Voltage-clamp recordings allow a clear separation of the initial rapid, AMPA receptor mediated excitatory postsynaptic current (EPSC) and the mGluR1-mediated slow EPSC (Fig. 1D) (Takechi et al. 1998; Hartmann et al. 2008). In addition of inducing the slow EPSPs, mGluR1s mediate a large and highly localized dendritic calcium transient in cerebellar Purkinje cells (Fig. 1D) (Llano et al. 1991a; Finch and Augustine 1998; Takechi et al. 1998).