P-Bodies and Stress Granules: Possible Roles in the Control of Translation and mRNA Degradation

The control of translation and mRNA degradation is important in the regulation of eukaryotic gene expression. In general, translation and steps in the major pathway of mRNA decay are in competition with each other. mRNAs that are not engaged in translation can aggregate into cytoplasmic mRNP granules referred to as processing bodies (P-bodies) and stress granules, which are related to mRNP particles that control translation in early development and neurons. Analyses of P-bodies and stress granules suggest a dynamic process, referred to as the mRNA Cycle, wherein mRNPs can move between polysomes, P-bodies and stress granules although the functional roles of mRNP assembly into higher order structures remain poorly understood. In this article, we review what is known about the coupling of translation and mRNA degradation, the properties of P-bodies and stress granules, and how assembly of mRNPs into larger structures might influence cellular function.The translation and decay of mRNAs play key roles in the control of eukaryotic gene expression. The determination of eukaryotic mRNA decay pathways has allowed insight into how translation and mRNA degradation are coupled. Degradation of eukaryotic mRNAs is generally initiated by shortening of the 3′ poly (A) tail (Fig. 1A) (reviewed in Parker and Song 2004; Garneau et al. 2007) by the major mRNA deadenylase, the Ccr4/Pop2/Not complex (Daugeron et al. 2001; Tucker et al. 2001; Thore et al. 2003). Following deadenylation, mRNAs can be degraded 3′ to 5′ by the exosome (Anderson and Parker 1998; Wang and Kiledjian 2001). However, more commonly, mRNAs are decapped by the Dcp1/Dcp2 decapping enzyme and then degraded 5′ to 3′ by the exonuclease, Xrn1 (Decker and Parker 1993; Hsu and Stevens 1993; Muhlrad et al. 1994, 1995; Dunckley and Parker 1999; van Dijk et al. 2002; Steiger et al. 2003). In metazoans, a second decapping enzyme, Nudt16, also contributes to mRNA turnover (Song et al. 2010).Open in a separate windowFigure 1.Eukaryotic mRNA decay pathways. (A) General mRNA decay pathways. (B) Specialized decay pathways that degrade translationally aberrant mRNAs.The processes of mRNA decay and translation are interconnected in eukaryotic cells in many ways. For example, quality control mechanisms exist to detect aberrancies in translation, which then lead to mRNAs being degraded by specialized mRNA decay pathways (Fig. 1B). Nonsense-mediated decay (NMD) is one such mRNA quality control system that degrades mRNAs that terminate translation aberrantly. In yeast, aberrant translation termination leads to deadenylation-independent decapping (Muhlrad and Parker 1994), whereas in metazoan cells NMD substrates can be both decapped and endonucleolytically cleaved and degraded (reviewed in Isken and Maquat 2007). A second quality control system for mRNA translation is referred to as no-go decay (NGD) and leads to endonucleolytic cleavage of mRNAs with strong stalls in translation elongation (Doma and Parker 2006; reviewed in Harigaya and Parker 2010). Another mechanism of mRNA quality control is the rapid 3′ to 5′ degradation of mRNAs that do not contain translation termination codons, which is referred to as non-stop decay (NSD) (Frischmeyer et al. 2002; van Hoof et al. 2002). The available evidence suggests these specialized mechanisms function primarily on aberrant mRNAs that are produced by defects in splicing, 3′ end formation, or damage to RNAs.The main pathway of mRNA degradation is also in competition with translation initiation. Competition between the two processes was first suggested by the observation that removal of the poly (A) tail and the cap structure, both of which stimulate translation initiation, were the key steps in mRNA degradation. In addition, inhibition of translation initiation by strong secondary structures in the 5′UTR, translation initiation inhibitors, a poor AUG context, or mutations in initiation factors increases the rates of deadenylation and decapping (Muhlrad et al. 1995; Muckenthaler et al. 1997; Lagrandeur and Parker 1999; Schwartz and Parker 1999). Moreover, the cap binding protein eIF4E, known to stimulate translation initiation, inhibits the decapping enzyme, Dcp1/Dcp2, both in vivo and in vitro (Schwartz and Parker 1999; Schwartz and Parker 2000). Finally, many mRNA specific regulatory factors, (e.g., miRNAs or PUF proteins), both repress translation and accelerate deadenylation and decapping (reviewed in Wickens et al. 2002; Behm-Ansmant et al. 2006; Franks and Lykke-Anderson 2008; Shyu et al. 2008).In the simplest model, the competition between translation and mRNA degradation can be understood through changes in the proteins bound to the cap and poly (A) tail that then influence the accessibility of these structures to deadenylases and decapping enzymes. For example, given that the Ccr4/Pop2/Not deadenylase complex is inhibited by poly (A)-binding protein (Pab1) (Tucker et al. 2002), the effects of translation on deadenylation are most likely through dynamic changes in the association of Pab1 binding with the poly (A) tail. One possibility is that defects in translation initiation either directly or indirectly decrease Pab1 association with the poly (A) tail. Deadenylation is also affected by aspects of translation termination. For instance, premature translation termination in yeast accelerates poly (A) shortening as part of the process of NMD (Cao and Parker 2003; Mitchell and Tollervey 2003). The coupling of translation termination to deadenylation has been suggested to occur through direct interactions of the translation termination factor eRF3 with Pab1 (Cosson et al. 2002), which may lead to Pab1 transiently dissociating from the poly (A) tail. Interestingly, in yeast, once the poly (A) tail reaches an oligo (A) length of 10–12 residues, a length that reduces the affinity of Pab1, the mRNA can become a substrate for decapping and for binding of the Pat1/Lsm1-7 complex (Tharun and Parker 2001; Chowdhury et al. 2007), which enhances the rate of decapping. This exchange of the Pab1 protein for the Pat1/Lsm1-7 complex is part of the mechanism that allows decapping to be promoted following deadenylation.A similar mRNP dynamic is also likely to occur on the cap structure. Specifically, the competition between translation initiation and decapping suggests that prior to decapping, translation initiation factors are exchanged for decapping factors, thereby assembling a distinct “decapping” mRNP that is no longer capable of translation initiation (Tharun and Parker 2001). This idea is supported by the observation that some decapping activators also function as translational repressors (Coller and Parker 2005; Pilkington and Parker 2008; Nissan et al. 2010). Thus, mRNA decapping appears to occur in two steps, first inhibition of translation initiation and exchange of translation factors for the general repression/degradation machinery, and a second step whereby the mRNA is actually degraded. Thus, by understanding the changes in mRNP states between actively translating mRNAs and mRNAs that are translationally repressed and possibly stored or ultimately degraded we will better understand how the fate of mRNAs is controlled in the cytoplasm.     

Protein Targeting and Transport as a Necessary Consequence of Increased Cellular Complexity

With increasing intracellular complexity, a new cell-biological problem that is the allocation of cytoplasmically synthesized proteins to their final destinations within the cell emerged. A special challenge is thereby the translocation of proteins into or across cellular membranes. The underlying mechanisms are only in parts well understood, but it can be assumed that the course of cellular evolution had a deep impact on the design of the required molecular machines. In this article, we aim to summarize the current knowledge and concepts of the evolutionary development of protein trafficking as a necessary premise and consequence of increased cellular complexity.

The evolution of modern cells is arguably the most challenging and important problem the field of biology has ever faced …—Carl R. Woese(Woese 2002)

Current models may accept that all modern eukaryotic cells arose from a single common ancestor (the cenancestral eukaryote), the nature of which is—owing to the lack of direct living or fossil descendants—still highly under debate (de Duve 2007). The chimeric nature of eukaryotic genomes with eubacterial and archaebacterial shares led to a discussion about the origin of this first “proto-eukaryote.” Several models exist (see Fig. 1), which either place the evolution of the nucleus before or after the emergence of the mitochondrion (outlined in Koonin 2010; Martijn and Ettema 2013). According to the different postulated scenarios (summarized in Embley and Martin 2006), eukaryotes in the latter case might have evolved by endosymbiosis between a hydrogen-producing, oxygen-producing, or sulfur-dependent α-proteobacterium and an archaebacterial host (Fig. 1C). The resulting mitochondriate prokaryote would have evolved the nucleus subsequently. In other scenarios (Fig. 1B), the cenancestral eukaryote emerged by cellular fusion or endosymbiosis of a Gram-negative, maybe hydrogen-producing, eubacterium and a methanogenic archaebacterium or eocyte, leading to a primitive but nucleated amitochondrial (archezoan) cell (Embley and Martin 2006, and references therein). As a third alternative, Cavalier-Smith (2002) suggested a common eubacterial ancestor for eukaryotes and archaebacteria (the Neomuran hypothesis) (Fig. 1A).Open in a separate windowFigure 1.Evolution of the last common ancestor of all eukaryotic cells. A schematic depiction of the early eukaryogenesis. Because of the lack of living and fossil descendants, several opposing models are discussed (A–C). The anticipated order of events is shown as a flow chart. For details, see text. (Derived from Embley and Martin 2006; Koonin 2010.)     

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.     

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

Receptor Tyrosine Kinases: Legacy of the First Two Decades

Receptor tyrosine kinases (RTKs) and their cellular signaling pathways play important roles in normal development and homeostasis. Aberrations in their activation or signaling leads to many pathologies, especially cancers, motivating the development of a variety of drugs that block RTK signaling that have been successfully applied for the treatment of many cancers. As the current field of RTKs and their signaling pathways are covered by a very large amount of literature, spread over half a century, I am focusing the scope of this review on seminal discoveries made before tyrosine phosphorylation was discovered, and on the early days of research into RTKs and their cellular signaling pathways. I review the history of the early days of research in the field of RTKs. I emphasize key early findings, which provided conceptual frameworks for addressing the questions of how RTKs are activated and how they regulate intracellular signaling pathways.The family of cell-surface receptors designated receptor tyrosine kinases (RTK) received their name more that a decade after the same molecules were already known as the cell-surface receptors for insulin (insulin receptor), epidermal growth factor (EGFR), and many other growth factor receptors. Following the pioneering discoveries of nerve growth factor and epidermal growth factor (EGF; Levi-Montalcini and Booker 1960; Cohen 1962) and the establishment of the important roles of these two growth factors in the control of neuronal differentiation and cell proliferation in vivo and in vitro, it became clear that these cytokines bind specifically to cell-surface receptors. Insulin had already been discovered by this time, and had been applied successfully to treat diabetes patients since the early twentieth century. The resulting homogenous preparations of pure insulin enabled the quantitative characterization of insulin binding to its receptor on intact cells or to solubilized insulin receptor preparations using radiolabeled insulin (De Meyts et al. 1973). These studies greatly advanced understanding of the ligand binding characteristics of insulin receptor and, later on EGFR (Carpenter et al. 1975), including the establishment of negative cooperativity in insulin binding to its receptor expressed on the surface of living cells (De Meyts et al. 1973). Moreover, these studies shed important light on the dynamic nature of the cellular behavior of these receptors. The capacities of insulin receptor and EGFR to undergo ligand-dependent down-regulation and desensitization through receptor-mediated internalization and degradation (Carpenter and Cohen 1976; Gordon et al. 1978; Schlessinger et al. 1978a,b; Carpentier et al. 1979; Haigler et al. 1979) were also established well before the realization that growth factors receptors are endowed with intrinsic protein tyrosine kinase activities (Fig. 1).Open in a separate windowFigure 1.A time line of key findings during the history of RTKs, with emphasis on findings and discoveries that produced the conceptual framework in the development of the RTK field and its application for cancer therapy. References for the key findings are also presented in the text (Lee et al. 1985; Libermann et al. 1985; Margolis et al. 1990; Bottaro et al. 1991; Bae et al. 2009).Progress was also made in elucidating the role of growth factors in normal embryonic development, wound healing, and pathological conditions such as cancer. Early studies in the 1960s and 1970s showed that growth factors play an important role in oncogenesis induced by retroviruses and in the proliferation of tumor-derived cancer cells. Pioneering studies performed by Howard Temin (1966, 1967) showed that cancer cells need less insulin and serum growth factors for cell proliferation compared with normal cells, suggesting that cancer cells produce and use their own growth factors and/or use cellular processes that in normal cells are regulated by exogenously supplied growth factors; both predictions were subsequently confirmed. A variety of new polypeptide growth factors that stimulate cell proliferation by binding to receptors at the cell surface were subsequently discovered. Those include a growth factor isolated from human platelets designated platelet-derived growth factor (PDGF; Antoniades et al. 1979; Heldin et al. 1979), a growth factor isolated from bovine brain designated fibroblast growth factor (FGF; Gospodarowicz et al. 1978), a growth factor isolated from rat platelets that stimulates the proliferation of mature hepatocytes, designated hepatocyte growth factor (HGF; Nakamura et al. 1986). In addition to EGF, another growth factor that binds selectively to cells expressing EGFR was isolated from virally and chemically transformed cells, suggesting that this growth factor—designated transforming growth factor α—may play a role in oncogenesis by an autocrine mechanism (Roberts et al. 1980, 1982). This discovery provided further support to the earlier finding that transformation by murine and feline sarcoma viruses selectively interferes with EGF binding to EGFR in transformed cells (Todaro et al. 1976). Together with many other studies published since the 1980s, this work showed that growth factors and their receptors play numerous important roles during development and in many normal cellular processes as well as in pathologies such as cancer, diabetes, atherosclerosis, severe bone disorders, and tumor angiogenesis.Visualization of dynamic cellular redistribution of ligand/receptor complexes, and rapid receptor-mediated internalization of growth factors such as insulin or EGF, led to the proposal that cell-surface receptors for these ligands may play a passive role in delivering them to intracellular compartments in which internalized EGF or insulin molecules exert their actions (Vigneri et al. 1978; Podlecki et al. 1986; Jiang and Schindler 1990). In other words, according to this hypothesis, the biological signals induced by insulin or EGF were thought to be mediated by binding of the ligands themselves to intracellular target(s) in the cytoplasm or nucleus, with the role of the cell-surface receptor being to act as a “carrier” that delivers them directly to these targets. An alternative hypothesis was that insulin or EGF activates their cognate receptors at the cell surface, which in turn stimulate the production of an intracellular second messenger molecule analogous to cAMP in signaling by the G-protein-activating β-adrenergic receptor. Indeed, several potential second messengers that are generated in cells on stimulation with insulin or other growth factors were proposed before (and even after) it became clear that insulin receptor, EGFR, and other RTKs are endowed with intrinsic tyrosine kinase activity (Larner et al. 1979; Das 1980; Saltiel and Cuatrecasas 1986).A demonstration that anti-insulin receptor antibodies from the serum of certain diabetic patients could mimic cellular responses of insulin (Flier et al. 1977; Van Obberghen et al. 1979) provided the first conclusive answer to the question of whether the biological activity of growth factors is mediated directly or indirectly through their membrane receptors. This experiment ruled out the possibility that insulin receptor functions as a passive carrier that delivers insulin to an intracellular target to induce cellular responses. Studies showing that intact, bivalent antibodies against the insulin receptor can activate its signaling, whereas monovalent Fab fragments of the same antibodies cannot further argued that ligand-induced receptor dimerization or stimulation of a particular arrangement between two receptor molecules in a dimer can activate the insulin receptor (Kahn et al. 1978).A similar conclusion was reached using certain monoclonal antibodies that bind to the extracellular region of EGFR and block ligand binding (Schreiber et al. 1981). Whereas intact antibodies were able to mimic EGF in stimulating a variety of EGF-like responses including cell proliferation, monovalent Fab fragments of the same monoclonal EGFR antibodies failed to do so—and acted instead as EGFR antagonists (Schreiber et al. 1981, 1983). These experiments provided strong evidence both that EGFR plays a crucial role in mediating EGF-induced cellular responses and that EGFR is activated by ligand-induced receptor dimerization (Schreiber 1981, 1983).     

The Role of Functional Prion-Like Proteins in the Persistence of Memory

Prions are a self-templating amyloidogenic state of normal cellular proteins, such as prion protein (PrP). They have been identified as the pathogenic agents, contributing to a number of diseases of the nervous system. However, the discovery that the neuronal RNA-binding protein, cytoplasmic polyadenylation element-binding protein (CPEB), has a prion-like state that is involved in the stabilization of memory raised the possibility that prion-like proteins can serve normal physiological functions in the nervous system. Here, we review recent experimental evidence of prion-like properties of neuronal CPEB in various organisms and propose a model of how the prion-like state may stabilize memory.Prions are proteinaceous infectious agents that were discovered in the 1980s by Stanley Prusiner while studying Creutzfeldt–Jakob disease (Prusiner 1982). Prusiner and colleagues showed them to be an amyloidogenic, self-perpetuating, forms of a normal cellular protein, termed prion protein or PrP. Prp in its self-perpetuating state kills cells. Prusiner and colleagues found that PrPs exist in at least two conformations: monomeric and aggregated (Fig. 1). The transition among these forms occurs spontaneously and only the aggregated conformation is pathogenic. Soon, PrPs were found to contribute to other neurodegenerative disorders in people, including kuru, transmissible spongiform encephalopathies, as well as bovine spongiform encephalopathy in cows (Prusiner 1994; Aguzzi and Weissmann 1998).Open in a separate windowFigure 1.Pathogenic prions exist in two states (soluble and aggregated and self-perpetuating). The conversion from the soluble to the aggregated form is spontaneous and the aggregated, self-perpetuating form is often toxic and kills the cell.There is now a growing consensus that similar prion-like, self-templating mechanisms underlie a variety of neurodegenerative disorders, including amyotrophic lateral sclerosis, Alzheimer’s disease, Parkinson’s disease, and Huntington’s disease (Polymenidou and Cleveland 2012).Not all prions, however, appear to be disease causing. Fungal prions, for instance, are nontoxic, and some may even be beneficial to the cells that harbor them (Wickner 1994; Shorter and Lindquist 2005; Crow and Li 2011). In 2003, Si and Kandel serendipitously discovered a prion-like protein in multicellular eukaryotes—the nervous system of the marine snail Aplysia—whose aggregated and self-perpetuating form contributes to the maintenance of long-term changes in synaptic efficacy. This functional prion-like protein differs from pathogenic prions in two important ways: (1) The conversion to the prion-like state is regulated by a physiological signal, and (2) the aggregated form has an identified physiological function (Fig. 2). Recent identification of new functional prion-like proteins in various organisms, including human, supports the idea that nonpathogenic prions may perform a wide range of biologically meaningful roles (Coustou et al. 1997; Eaglestone et al. 1999; True and Lindquist 2000; Ishimaru et al. 2003; True et al. 2004; Hou et al. 2011; Jarosz et al. 2014).Open in a separate windowFigure 2.“Functional” prion: memory. “Functional” prions differ from conventional prions in two ways. First, the conversion is triggered by a physiological signal, and second, the aggregated, self-perpetrating forms have a physiological function. 5-HT, Serotonin; DA, dopamine.In this review, we focus on functional prion-like proteins in the brain and specifically on the prion-like properties of the cytoplasmic polyadenylation element-binding protein (CPEB), and examine how the prion-like state can control protein synthesis at the synapse and, thereby, synaptic plasticity and long-lasting memory. We anticipate the studies of CPEB would also provide some generalizable concepts as to how prion-based protein switches in multicellular eukaryotes may work.     

Base Excision Repair

Base excision repair (BER) corrects DNA damage from oxidation, deamination and alkylation. Such base lesions cause little distortion to the DNA helix structure. BER is initiated by a DNA glycosylase that recognizes and removes the damaged base, leaving an abasic site that is further processed by short-patch repair or long-patch repair that largely uses different proteins to complete BER. At least 11 distinct mammalian DNA glycosylases are known, each recognizing a few related lesions, frequently with some overlap in specificities. Impressively, the damaged bases are rapidly identified in a vast excess of normal bases, without a supply of energy. BER protects against cancer, aging, and neurodegeneration and takes place both in nuclei and mitochondria. More recently, an important role of uracil-DNA glycosylase UNG2 in adaptive immunity was revealed. Furthermore, other DNA glycosylases may have important roles in epigenetics, thus expanding the repertoire of BER proteins.Base excision repair (BER) corrects small base lesions that do not significantly distort the DNA helix structure. Such damage typically results from deamination, oxidation, or methylation (Fig. 1). Much of the damage is the result of spontaneous decay of DNA (Lindahl 1993), although similar damage may also be caused by environmental chemicals, radiation, or treatment with cytostatic drugs. BER takes place in nuclei, as well as in mitochondria, largely using different isoforms of proteins or genetically distant proteins. The identification of Escherichia coli uracil-DNA glycosylase (Ung) in 1974 by Tomas Lindahl marks the discovery of BER. Lindahl searched for an enzyme activity that would act on genomic uracil resulting from cytosine deamination. Such an activity was found, but rather unexpectedly, it was not a nuclease. Instead, Lindahl identified an enzyme that cleaved the bond between uracil and deoxyribose. The resulting abasic site (AP-site) was suggested to be further processed by an AP-endonuclease, an exonuclease, a DNA polymerase, and a ligase. Thus, the fundamental steps in the BER pathway were outlined already in the very first paper (Lindahl 1974). Enzymes that cleave the bond between deoxyribose and a modified or mismatched DNA base are now called DNA glycosylases. Collectively these enzymes initiate base excision repair of a large number of base lesions, each recognized by one or a few DNA glycosylases with overlapping specificities.Open in a separate windowFigure 1.Chemistry of common base lesions and abasic sites.This relatively brief review focuses on recent advances in the mechanism and function of BER with a focus on mammalian proteins. The current view is that BER is important in relation to cancer, neurodegeneration, and aging (Jeppesen et al. 2011; Wallace et al. 2012). Because of limited space, we have referred to reviews for the majority of results published more than 6–7 years ago. Also, for more detailed analyses of different aspects of BER, the reader is referred to excellent reviews on BER proteins and pathways published in Huffman et al. (2005), Beard and Wilson (2006), Berti and McCann (2006), Cortázar et al. (2007), Kavli et al. (2007), Sousa et al. (2007), Tubbs et al. (2007), Berger et al. (2008), Robertson et al. (2009), Friedman and Stivers (2010), Wilson et al. (2010), Svilar et al. (2011), and Jacobs and Schar (2012).     

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).     

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).     

The Structure and Function of the Eukaryotic Ribosome

Structures of the bacterial ribosome have provided a framework for understanding universal mechanisms of protein synthesis. However, the eukaryotic ribosome is much larger than it is in bacteria, and its activity is fundamentally different in many key ways. Recent cryo-electron microscopy reconstructions and X-ray crystal structures of eukaryotic ribosomes and ribosomal subunits now provide an unprecedented opportunity to explore mechanisms of eukaryotic translation and its regulation in atomic detail. This review describes the X-ray crystal structures of the Tetrahymena thermophila 40S and 60S subunits and the Saccharomyces cerevisiae 80S ribosome, as well as cryo-electron microscopy reconstructions of translating yeast and plant 80S ribosomes. Mechanistic questions about translation in eukaryotes that will require additional structural insights to be resolved are also presented.All ribosomes are composed of two subunits, both of which are built from RNA and protein (Figs. ​(Figs.11 and ​and2).2). Bacterial ribosomes, for example of Escherichia coli, contain a small subunit (SSU) composed of one 16S ribosomal RNA (rRNA) and 21 ribosomal proteins (r-proteins) (Figs. ​(Figs.1A1A and ​and1B)1B) and a large subunit (LSU) containing 5S and 23S rRNAs and 33 r-proteins (Fig. 2A). Crystal structures of prokaryotic ribosomal particles, namely, the Thermus thermophilus SSU (Schluenzen et al. 2000; Wimberly et al. 2000), Haloarcula marismortui and Deinococcus radiodurans LSU (Ban et al. 2000; Harms et al. 2001), and E. coli and T. thermophilus 70S ribosomes (Yusupov et al. 2001; Schuwirth et al. 2005; Selmer et al. 2006), reveal the complex architecture that derives from the network of interactions connecting the individual r-proteins with each other and with the rRNAs (Brodersen et al. 2002; Klein et al. 2004). The 16S rRNA can be divided into four domains, which together with the r-proteins constitute the structural landmarks of the SSU (Wimberly et al. 2000) (Fig. 1A): The 5′ and 3′ minor (h44) domains with proteins S4, S5, S12, S16, S17, and S20 constitute the body (and spur or foot) of the SSU; the 3′ major domain forms the head, which is protein rich, containing S2, S3, S7, S9, S10, S13, S14, and S19; whereas the central domain makes up the platform by interacting with proteins S1, S6, S8, S11, S15, and S18 (Fig. 1B). The rRNA of the LSU can be divided into seven domains (including the 5S rRNA as domain VII), which—in contrast to the SSU—are intricately interwoven with the r-proteins as well as each other (Ban et al. 2000; Brodersen et al. 2002) (Fig. 2A). Structural landmarks on the LSU include the central protuberance (CP) and the flexible L1 and L7/L12 stalks (Fig. 2A).Open in a separate windowFigure 1.The bacterial and eukaryotic small ribosomal subunit. (A,B) Interface (upper) and solvent (lower) views of the bacterial 30S subunit (Jenner et al. 2010a). (A) 16S rRNA domains and associated r-proteins colored distinctly: b, body (blue); h, head (red); pt, platform (green); and h44, helix 44 (yellow). (B) 16S rRNA colored gray and r-proteins colored distinctly and labeled. (C–E) Interface and solvent views of the eukaryotic 40S subunit (Rabl et al. 2011), with (C) eukaryotic-specific r-proteins (red) and rRNA (pink) shown relative to conserved rRNA (gray) and r-proteins (blue), and with (D,E) 18S rRNA colored gray and r-proteins colored distinctly and labeled.Open in a separate windowFigure 2.The bacterial and eukaryotic large ribosomal subunit. (A) Interface (upper) and solvent (lower) views of the bacterial 50S subunit (Jenner et al. 2010b), with 23S rRNA domains and bacterial-specific (light blue) and conserved (blue) r-proteins colored distinctly: cp, central protuberance; L1, L1 stalk; and St, L7/L12 stalk (or P-stalk in archeaa/eukaryotes). (B–E) Interface and solvent views of the eukaryotic 60S subunit (Klinge et al. 2011), with (B) eukaryotic-specific r-proteins (red) and rRNA (pink) shown relative to conserved rRNA (gray) and r-proteins (blue), (C) eukaryotic-specific expansion segments (ES) colored distinctly, and (D,E) 28S rRNA colored gray and r-proteins colored distinctly and labeled.In contrast to their bacterial counterparts, eukaryotic ribosomes are much larger and more complex, containing additional rRNA in the form of so-called expansion segments (ES) as well as many additional r-proteins and r-protein extensions (Figs. 1C–E and ​and2C–E).2C–E). Compared with the ∼4500 nucleotides of rRNA and 54 r-proteins of the bacterial 70S ribosome, eukaryotic 80S ribosomes contain >5500 nucleotides of rRNA (SSU, 18S rRNA; LSU, 5S, 5.8S, and 25S rRNA) and 80 (79 in yeast) r-proteins. The first structural models for the eukaryotic (yeast) ribosome were built using 15-Å cryo–electon microscopy (cryo-EM) maps fitted with structures of the bacterial SSU (Wimberly et al. 2000) and archaeal LSU (Ban et al. 2000), thus identifying the location of a total of 46 eukaryotic r-proteins with bacterial and/or archaeal homologs as well as many ES (Spahn et al. 2001a). Subsequent cryo-EM reconstructions led to the localization of additional eukaryotic r-proteins, RACK1 (Sengupta et al. 2004) and S19e (Taylor et al. 2009) on the SSU and L30e (Halic et al. 2005) on the LSU, as well as more complete models of the rRNA derived from cryo-EM maps of canine and fungal 80S ribosomes at ∼9 Å (Chandramouli et al. 2008; Taylor et al. 2009). Recent cryo-EM reconstructions of plant and yeast 80S translating ribosomes at 5.5–6.1 Å enabled the correct placement of an additional six and 10 r-proteins on the SSU and LSU, respectively, as well as the tracing of many eukaryotic-specific r-protein extensions (Armache et al. 2010a,b). The full assignment of the r-proteins in the yeast and fungal 80S ribosomes, however, only became possible with the improved resolution (3.0–3.9 Å) resulting from the crystal structures of the SSU and LSU from Tetrahymena thermophila (Klinge et al. 2011; Rabl et al. 2011) and the Saccharomyces cerevisiae 80S ribosome (Figs. ​(Figs.1D,E1D,E and ​and2D,E)2D,E) (Ben-Shem et al. 2011).     

The Desmosome

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