Schwann Cells: Development and Role in Nerve Repair

Schwann cells develop from the neural crest in a well-defined sequence of events. This involves the formation of the Schwann cell precursor and immature Schwann cells, followed by the generation of the myelin and nonmyelin (Remak) cells of mature nerves. This review describes the signals that control the embryonic phase of this process and the organogenesis of peripheral nerves. We also discuss the phenotypic plasticity retained by mature Schwann cells, and explain why this unusual feature is central to the striking regenerative potential of the peripheral nervous system (PNS).The myelin and nonmyelin (Remak) Schwann cells of adult nerves originate from the neural crest in well-defined developmental steps (Fig. 1). This review focuses on embryonic development (for additional information on myelination, see Salzer 2015). We also discuss how the ability to change between differentiation states, a characteristic attribute of developing cells, is retained by mature Schwann cells, and explain how the ability of Schwann cells to change phenotype in response to injury allows the peripheral nervous system (PNS) to regenerate after damage.Open in a separate windowFigure 1.Main transitions in the Schwann cell precursor (SCP) lineage. The diagram shows both developmental and injury-induced transitions. Black uninterrupted arrows, normal development; red arrows, the Schwann cell injury response; stippled arrows, postrepair reformation of myelin and Remak cells. Embryonic dates (E) refer to mouse development. (Modified from Jessen and Mirsky 2012; reprinted, with permission and with contribution from Y. Poitelon and L. Feltri.)     

Polarity in Stem Cell Division: Asymmetric Stem Cell Division in Tissue Homeostasis

Many adult stem cells divide asymmetrically to balance self-renewal and differentiation, thereby maintaining tissue homeostasis. Asymmetric stem cell divisions depend on asymmetric cell architecture (i.e., cell polarity) within the cell and/or the cellular environment. In particular, as residents of the tissues they sustain, stem cells are inevitably placed in the context of the tissue architecture. Indeed, many stem cells are polarized within their microenvironment, or the stem cell niche, and their asymmetric division relies on their relationship with the microenvironment. Here, we review asymmetric stem cell divisions in the context of the stem cell niche with a focus on Drosophila germ line stem cells, where the nature of niche-dependent asymmetric stem cell division is well characterized.Asymmetric cell division allows stem cells to self-renew and produce another cell that undergoes differentiation, thus providing a simple method for tissue homeostasis. Stem cell self-renewal refers to the daughter(s) of stem cell division maintaining all stem cell characteristics, including proliferation capacity, maintenance of the undifferentiated state, and the capability to produce daughter cells that undergo differentiation. A failure to maintain the correct stem cell number has been speculated to lead to tumorigenesis/tissue hyperplasia via stem cell hyperproliferation or tissue degeneration/aging via a reduction in stem cell number or activity (Morrison and Kimble 2006; Rando 2006). This necessity changes during development. The stem cell pool requires expansion earlier in development, whereas maintenance is needed later to sustain tissue homeostasis.There are two major mechanisms to sustain a fixed number of adult stem cells: stem cell niche and asymmetric stem cell division, which are not mutually exclusive. Stem cell niche is a microenvironment in which stem cells reside, and provides essential signals required for stem cell identity (Fig. 1A). Physical limitation of niche “space” can therefore define stem cell number within a tissue. Within such a niche, many stem cells divide asymmetrically, giving rise to one stem cell and one differentiating cell, by placing one daughter inside and another outside of the niche, respectively (Fig. 1A). Nevertheless, some stem cells divide asymmetrically, apparently without the niche. For example, in Drosophila neuroblasts, cell-intrinsic fate determinants are polarized within a dividing cell, and subsequent partitioning of such fate determinants into daughter cells in an asymmetric manner results in asymmetric stem cell division (Fig. 1B) (see Fig. 3A and Prehoda 2009).Open in a separate windowFigure 1.Mechanisms of asymmetric stem cell division. (A) Asymmetric stem cell division by extrinsic fate determinants (i.e., the stem cell niche). The two daughters of stem cell division will be placed in distinct cellular environments either inside or outside the stem cell niche, leading to asymmetric fate choice. (B) Asymmetric stem cell division by intrinsic fate determinants. Fate determinants are polarized in the dividing stem cells, which are subsequently partitioned into two daughter cells unequally, thus making the division asymmetrical. Self-renewing (red line) and/or differentiation promoting (green line) factors may be involved.In this review, we focus primarily on asymmetric stem cell divisions in the Drosophila germ line as the most intensively studied examples of niche-dependent asymmetric stem cell division. We also discuss some examples of stem cell division outside Drosophila, where stem cells are known to divide asymmetrically or in a niche-dependent manner.     

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).     

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.)     

Mitochondrial Regulation of Cell Death

Although required for life, paradoxically, mitochondria are often essential for initiating apoptotic cell death. Mitochondria regulate caspase activation and cell death through an event termed mitochondrial outer membrane permeabilization (MOMP); this leads to the release of various mitochondrial intermembrane space proteins that activate caspases, resulting in apoptosis. MOMP is often considered a point of no return because it typically leads to cell death, even in the absence of caspase activity. Because of this pivotal role in deciding cell fate, deregulation of MOMP impacts on many diseases and represents a fruitful site for therapeutic intervention. Here we discuss the mechanisms underlying mitochondrial permeabilization and how this key event leads to cell death through caspase-dependent and -independent means. We then proceed to explore how the release of mitochondrial proteins may be regulated following MOMP. Finally, we discuss mechanisms that enable cells sometimes to survive MOMP, allowing them, in essence, to return from the point of no return.In most organisms, mitochondria play an essential role in activating caspase proteases through a pathway termed the mitochondrial or intrinsic pathway of apoptosis. Mitochondria regulate caspase activation by a process called mitochondrial outer membrane permeabilization (MOMP). Selective permeabilization of the mitochondrial outer membrane releases intermembrane space (IMS) proteins that drive robust caspase activity leading to rapid cell death. However, even in the absence of caspase activity, MOMP typically commits a cell to death and is therefore considered a point of no return (Fig. 1). Because of this pivotal role in dictating cell fate, MOMP is highly regulated, mainly through interactions between pro- and antiapoptotic members of the Bcl-2 family. In this article, we begin by discussing how mitochondria may have evolved to become central players in apoptotic cell death. We then provide an overview of current models addressing the mechanics of MOMP, outlining how this crucial event leads to cell death through both caspase-dependent or -independent mechanisms. Finally, we discuss how caspase activity may be regulated post-MOMP and define other processes that allow cells to survive MOMP and, in effect, return from the point of no return.Open in a separate windowFigure 1.Mitochondrial regulation of cell death. Bax/Bak-mediated mitochondrial outer membrane permeabilization (MOMP) can lead to caspase-dependent apoptosis (left) or caspase-independent cell death (right). Following MOMP, soluble proteins are released from the mitochondrial intermembrane space into the cytoplasm. Cytochrome c binds to monomeric Apaf-1 leading to its conformational change and oligomerization. Procaspase-9 is recruited to heptameric Apaf-1 complexes forming the apoptosome. This leads to activation of caspase-9 and, through caspase-9-mediated cleavage, activation of the executioner caspases-3 and -7. Release of Smac and Omi from the mitochondrial intermembrane space facilitates caspase activation by neutralizing the caspase inhibitor XIAP. MOMP can also lead to nonapoptotic cell death through a gradual loss of mitochondrial function and/or release of mitochondrial proteins that kill the cell in a caspase-independent manner.     

Cyanobacterial Heterocysts

Many multicellular cyanobacteria produce specialized nitrogen-fixing heterocysts. During diazotrophic growth of the model organism Anabaena (Nostoc) sp. strain PCC 7120, a regulated developmental pattern of single heterocysts separated by about 10 to 20 photosynthetic vegetative cells is maintained along filaments. Heterocyst structure and metabolic activity function together to accommodate the oxygen-sensitive process of nitrogen fixation. This article focuses on recent research on heterocyst development, including morphogenesis, transport of molecules between cells in a filament, differential gene expression, and pattern formation.Organisms composed of multiple differentiated cell types can possess structures, functions, and behaviors that are more diverse and efficient than those of unicellular organisms. Among multicellular prokaryotes, heterocyst-forming cyanobacteria offer an excellent model for the study of cellular differentiation and multicellular pattern formation. Cyanobacteria are a large group of Gram-negative prokaryotes that perform oxygenic photosynthesis. They have evolved multiple specialized cell types, including nitrogen-fixing heterocysts, spore-like akinetes, and the cells of motile hormogonia filaments. Of these, the development of heterocysts in the filamentous cyanobacterium Anabaena (also Nostoc) sp. strain PCC 7120 (hereafter Anabaena PCC 7120) has been the best studied. Heterocyst development offers a striking example of cellular differentiation and developmental biology in a very simple form: Filaments are composed of only two cell types and these are arrayed in a one-dimensional pattern similar to beads on a string (Figs. 1 and ​and22).Open in a separate windowFigure 1.Heterocyst development in Anabaena PCC 7120. (A) Anabaena PCC 7120 grown in medium containing a source of combined nitrogen grows as filaments of photosynthetic vegetative cells. (B) In the absence of combined nitrogen, heterocysts differentiate at semiregular intervals, forming a developmental pattern of single heterocysts every 10 to 20 vegetative cells along filaments. Heterocysts are often larger than vegetative cells, have a thicker multilayered envelope, and usually contain cyanophycin granules at their poles adjacent to a vegetative cell.Open in a separate windowFigure 2.Heterocyst development in Anabaena PCC 7120. Filaments of the wild type carrying a patS-gfp reporter grown in medium containing nitrate are composed of vegetative cells (A), and have undergone heterocyst development 1 d after transfer to medium without combined nitrogen (B). A patS mutant strain carrying the same patS-gfp reporter grown in media containing nitrate contains a small number of heterocysts (C), and 1 d after transfer to medium without combined nitrogen shows a higher than normal frequency of heterocysts and an abnormal developmental pattern (D). (A, B, C, D) Merged DIC (grayscale), autofluorescence of photosynthetic pigments (red), and patS-gfp reporter fluorescence (green) microscopic images; arrowheads indicate heterocysts; asterisks indicate proheterocysts; size bar, 5 µm. (E, F) Transmission electron micrographs of wild-type vegetative cells (V) and a heterocyst (H) at the end of a filament; T, thylakoid membranes; PS, polysaccharide layer; GL, glycolipid layer; C, polar cyanophycin granule; size bar, 0.2 µm.Many cyanobacterial species are capable of nitrogen fixation. However, oxygenic photosynthesis and nitrogen fixation are incompatible processes because nitrogenase is inactivated by oxygen. Cyanobacteria mainly use two mechanisms to separate these activities: a biological circadian clock to separate them temporally, and multicellularity and cellular differentiation to separate them spatially. For example, the unicellular Cyanothece sp. strain ATCC 51142 stores glycogen during the day and fixes nitrogen at night (Toepel et al. 2008), whereas the filamentous Trichodesmium erythraeum IMS101 fixes nitrogen during the day in groups of specialized cells (Sandh et al. 2009). Heterocyst-forming cyanobacteria differentiate highly specialized cells to provide fixed nitrogen to the vegetative cells in a filament.In the presence of a source of combined nitrogen such as nitrate or ammonium, Anabaena PCC 7120 grows as long filaments containing hundreds of photosynthetic vegetative cells. In the absence of combined nitrogen, it produces heterocysts, which are terminally differentiated nitrogen-fixing cells that form at semiregular intervals between stretches of vegetative cells to produce a multicellular pattern of single heterocysts every ten to twenty vegetative cells along filaments (Figs. 1 and ​and2).2). Some heterocyst-forming cyanobacteria show different regulation or display different developmental patterns but these topics are beyond the scope of this article. Heterocyst development involves integration of multiple external and internal signals, communication between the cells in a filament, and temporal and spatial regulation of genes and cellular processes. The study of heterocyst development in Anabaena PCC 7120 has proven to be an excellent model for the study of cell fate determination, pattern formation, and differential gene expression during prokaryotic multicellular evelopment. Various aspects of heterocyst development, signaling, and regulation have been the subject of several recent reviews (Meeks and Elhai 2002; Forchhammer 2004; Herrero et al. 2004; Zhang et al. 2006; Aldea et al. 2008; Zhao and Wolk 2008).Although beyond the scope of this article, it should be noted that cyanobacteria have recently attracted increased attention because of their important roles in environmental carbon and nitrogen fixation (Montoya et al. 2004), and their potential for providing renewable chemicals and biofuels (Dismukes et al. 2008).     

Forming Patterns in Development without Morphogen Gradients: Scattered Differentiation and Sorting Out

Few mechanisms provide alternatives to morphogen gradients for producing spatial patterns of cells in development. One possibility is based on the sorting out of cells that initially differentiate in a salt and pepper mixture and then physically move to create coherent tissues. Here, we describe the evidence suggesting this is the major mode of patterning in Dictyostelium. In addition, we discuss whether convergent evolution could have produced a conceptually similar mechanism in other organisms.A limited number of processes are thought to regulate the differentiation of specialized cell types and their organization to form larger scale structures, such as organs or limbs, during embryonic development. First, early embryological experiments revealed a patterning process that depends on special “organizing” regions in the embryo. This idea was encapsulated as “positional information” and led to the concept of morphogen gradients (Fig. 1) (Wolpert 1996). In addition, cytoplasmic determinants have been shown to direct development along different lines when they are partitioned unequally between daughter cells by asymmetric cell division (Betschinger and Knoblich 2004). Finally, short-range inductive signaling can specify cells at a local level and when reiterated produces highly ordered structures (Simpson 1990; Freeman 1997; Meinhardt and Gierer 2000).Open in a separate windowFigure 1.Alternative ways of patterning cells during development. (A) Patterning by “positional information”: A group of undifferentiated cells is patterned by a morphogen diffusing from a pre-established source, producing a concentration gradient. Cells respond according to the local morphogen concentration, becoming red, white, or blue. (B, C) Patterning without positional information: This is a two-step process in which different cell types first differentiate mixed up with each other, and then sort out. The initial differentiation can be controlled by strictly local interactions between the cells, as in lateral inhibition (B), or by a global signal to which cells respond with different sensitivities and whose concentration they regulate by negative feedback (C). Once sorting has occurred, the global inducer forms a reverse gradient, which could then convey positional information for further patterning events.The question then arises of whether evolution has devised any further global patterning mechanisms. One possibility that has been repeatedly considered, but not firmly established as a general mechanism, is based on sorting out. In this process, pattern is produced in two steps: (1) Different cell types are initially specified from a precursor pool independent of their position to produce a salt and pepper mixture and (2) the mixture of cell types is resolved into discrete tissues by the physical movement and sorting out of the cells (Fig. 1). Consequently, this mechanism does not involve positional information. However, it can actually provide the conditions under which positional signaling and morphogen gradients can arise, if the resolved tissues then act as sources and sinks for signal molecules.We first describe the powerful evidence that this alternative patterning process is used during the developmental cycle of the social amoeba Dictyostelium discoideum, and then consider the possibility that this patterning strategy may be used more widely.     

The Signaling Mechanisms Underlying Cell Polarity and Chemotaxis

Chemotaxis—the directed movement of cells in a gradient of chemoattractant—is essential for neutrophils to crawl to sites of inflammation and infection and for Dictyostelium discoideum (D. discoideum) to aggregate during morphogenesis. Chemoattractant-induced activation of spatially localized cellular signals causes cells to polarize and move toward the highest concentration of the chemoattractant. Extensive studies have been devoted to achieving a better understanding of the mechanism(s) used by a neutrophil to choose its direction of polarity and to crawl effectively in response to chemoattractant gradients. Recent technological advances are beginning to reveal many fascinating details of the intracellular signaling components that spatially direct the cytoskeleton of neutrophils and D. discoideum and the complementary mechanisms that make the cell''s front distinct from its back.Chemotaxis—the directed movement of cells in a gradient of chemoattractant—allows leukocytes to seek out sites of inflammation and infection, amoebas of Dictyostelium discoideum (D. discoideum) to aggregate, neurons to send projections to specific regions of the brain to find their synaptic partners, yeast cells to mate, and fibroblasts to move into the wound space (Fig. 1). In each case, chemoattractant-induced activation of spatially localized cellular signals causes cells to polarize and move toward the highest concentration of the chemoattractant. During chemotaxis, filamentous actin (F-actin) is polymerized asymmetrically at the upgradient edge of the cell (leading edge), providing the necessary force to thrust projections of the plasma membrane in the proper direction (see Mullins 2009). Neutrophilic leukocytes (neutrophils), for instance, can polarize and move up very shallow gradients, with a chemoattractant concentration ∼2% higher at the front than the back (Fig. 2) (Devreotes and Zigmond 1988). To restrict actin polymerization to the leading edge in such a shallow gradient, neutrophils must create a much steeper internal gradient of regulatory signals. In addition, distinctive actin–myosin contractile complexes are also formed at the sides and back of the cells (Fig. 2). The ability to create such distinctive segregation of actin assemblies enables neutrophils to move nearly 50 times more quickly than fibroblasts. The polarization response is self-organizing, which occurs even when the attractant concentration is uniform and apparently stimulating all portions of the plasma membrane at the same intensity; in the absence of a gradient, the direction of polarity is random, but all cells can be induced to polarize (Fig. 2). Thus, neutrophil polarization to chemoattractant stimulation represents a striking example of symmetry breaking from an unpolarized state to a polarized one.Open in a separate windowFigure 1.Examples of chemotaxis. (A) A human neutrophil chasing a Staphylococcus aureus microorganism on a blood film among red blood cells, notable for their dark color and principally spherical shape (imaged by David Rogers, courtesy of Thomas P. Stossel). Bar, 10 µm. Chemotaxis is also necessary for (B) D. discoideum to form multicellular aggregates during development (courtesy of M.J. Grimson and R.L. Blanton, Texas Tech University), and (C) for axons to find their way in the developing nervous system. Photo provided by Kathryn Tosney, University of Miami.Open in a separate windowFigure 2.(A–D) Polarization of a neutrophil in response to gradient of chemoattractant. Nomarski images of unpolarized neutrophil responding to a micropipette containing the chemoattractant fMLP (white circle) at (A) 5 s, (B) 30 s, (C) 81 s, and (D) 129 s of stimulation. Bar = 5 µm. (Figure is taken from Weiner et al. 1999, with permission.) Human neutrophils stimulated with fMLP show highly polarized morphology and asymmetric cytoskeletal assemblies. (E–G) Human neutrophils were stimulated by a uniform concentration of fMLP (100 nM) and fixed 2 min after stimulations. Fixed cells were stained for F-actin with rhodamine-phalloidin (E, red) and an antibody raised against activated myosin II (phosphorylated specifically at Ser19, p[19]-MLC) (F, green). These fluorescent images are merged with Nomarski image in (G). Bars, 10 µm.To enter an infected tissue, neutrophils require chemoattractants produced by host cells and microorganisms to migrate to the sites and infection and inflammation. Neutrophil chemotaxis also contributes to many inflammatory and autoimmune diseases, including rheumatoid arthritis, ischemia-reperfusion syndrome, acute respiratory distress, and systemic inflammatory response syndromes. Although the critical physiological functions of neutrophils have made their chemoattractants and chemoattractant receptors targets of intense investigation, understanding of the neutrophil polarity and directional migration has until recently lagged behind that of other cells. Over the past decade, experimentation with knockout mice and human neutrophil cell lines has begun to shed light on the complex intracellular signals responsible for neutrophil polarity. In this article, I summarize recent advances in the study of chemotactic signals in neutrophils, with some of the discussion also devoted to a related model—chemotaxis of D. discoideum. These soil amoebas grow as single cells, but on starvation chemotax into multicellular aggregates in response to secreted chemoattractants such as adenosine 3′,5′-monophosphate (cAMP).     

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).     

Golgi Glycosylation

Glycosylation is a very common modification of protein and lipid, and most glycosylation reactions occur in the Golgi. Although the transfer of initial sugar(s) to glycoproteins or glycolipids occurs in the ER or on the ER membrane, the subsequent addition of the many different sugars that make up a mature glycan is accomplished in the Golgi. Golgi membranes are studded with glycosyltransferases, glycosidases, and nucleotide sugar transporters arrayed in a generally ordered manner from the cis-Golgi to the trans-Golgi network (TGN), such that each activity is able to act on specific substrate(s) generated earlier in the pathway. The spectrum of glycosyltransferases and other activities that effect glycosylation may vary with cell type, and thus the final complement of glycans on glycoconjugates is variable. In addition, glycan synthesis is affected by Golgi pH, the integrity of Golgi peripheral membrane proteins, growth factor signaling, Golgi membrane dynamics, and cellular stress. Knowledge of Golgi glycosylation has fostered the development of assays to identify mechanisms of intracellular vesicular trafficking and facilitated glycosylation engineering of recombinant glycoproteins.The Golgi is home to a multitude of glycosyltransferases (GTs), glycosidases, and nucleotide sugar transporters that function together to complete the synthesis of glycans from founding sugars covalently attached to protein or lipid in the endoplasmic reticulum (ER) (Fig. 1, sugars shaded in green). Thus, glycoproteins, glycosphingolipids (GSLs), proteoglycans, and glycophosphatidylinositol (GPI) anchors acquire their final sugar complement during passage through the Golgi. Most glycoproteins and proteoglycans are either secreted from the cell, or span the plasma membrane with their glycans becoming the molecular frontier of the cell (Fig. 1). GSLs and GPI-anchored proteins also reside in the plasma membrane, the latter being confined to the outer leaflet of the lipid bilayer. The forest of glycans at the cell surface is often called the glycocalyx and can be visualized by electron microscopy after staining for sugars.Open in a separate windowFigure 1.Glycans that mature in the Golgi. The diagram depicts simple N- and O-glycans attached to glycoproteins, proteoglycans, glycosphingolipids, and a GPI anchor in the plasma membrane. Rather rare O-glycans are found attached to EGF-like repeats (EGF; pink) or thrombospondin repeats (TSR; gray) with a particular consensus sequence. The WxxW motif in a TSR is C-mannosylated. Core regions boxed in teal are sugars added in the ER. The remaining sugars in each class of glycan are added during passage through the cis-, medial-, and trans-Golgi network (TGN) compartments of the Golgi. Abbreviations are: Man, mannose; Gal, galactose; Glc, glucose; GlcNAc, N-acetylglucosamine; GlcNH₂, Glucosamine; GlcA, glucuronic acid; IdoA, iduronic acid; GalNAc, N-acetylgalactosamine; Xyl, xylose; Fuc, Fucose; Sia, sialic acid; 3S, 3-O-sulfated; 6S, 6-O-sulfated, PO₄⁻, phosphate. (Modified from Figure 1.6 in Essentials of glycobiology, with permission from Varki and Sharon 2009.)Glycosylation is the most common posttranslational modification of proteins. Mature glycans at any one glycosylation site may be as simple as a single sugar, or as complex as a polymer of more than 200 sugars, potentially modified with phosphate, sulfate, acetate, or phosphorylcholine. Most importantly, glycans are often branched. For example, a complex N-glycan (Fig. 1) may have up to six branches or antennae, and each antenna may contain many repeating disaccharide units. This article will describe the nature of resident Golgi GTs and other activities involved in Golgi glycosylation from entry into the cis-Golgi through passage to the trans-Golgi network (TGN). The focus is on mammalian Golgi glycosylation but comparisons with yeast, Caenorhabditis elegans, and Drosophila are made where appropriate.     

Physical Model of Cellular Symmetry Breaking

Cells can polarize in response to external signals, such as chemical gradients, cell–cell contacts, and electromagnetic fields. However, cells can also polarize in the absence of an external cue. For example, a motile cell, which initially has a more or less round shape, can lose its symmetry spontaneously even in a homogeneous environment and start moving in random directions. One of the principal determinants of cell polarity is the cortical actin network that underlies the plasma membrane. Tension in this network generated by myosin motors can be relaxed by rupture of the shell, leading to polarization. In this article, we discuss how simplified model systems can help us to understand the physics that underlie the mechanics of symmetry breaking.Symmetry breaking in physics is an old well-known concept. It is based on energy considerations: A symmetrical system can lose its symmetry if an asymmetrical state has a lower energy. The initial symmetrical state can be either unstable or metastable. In the latter case, there is an energy barrier to be overcome before symmetry breaking occurs. An external trigger can drive the system from its symmetrical to its asymmetrical state, but simple noise can also do so if its amplitude is sufficiently high. A simple example is a clown balancing on a ball: When the clown is standing on top of the ball, the system has a cylindrical symmetry (Fig. 1A). However, this state is unstable: The slightest perturbation will cause the clown to fall down in some direction, breaking the cylindrical symmetry (Fig. 1B). Imagine now that the ball is slightly flat on its base, giving more stability to the clown. Such a state is metastable: The clown can make small excursions safely (Fig. 1C,D), but if he moves too much (i.e., generates too much “noise”), he will fall down in this case also (Fig. 1E,F).Open in a separate windowFigure 1.Illustration of symmetry breaking with a clown standing on a balloon. In (A), the clown is in unstable equilibrium and the situation is symmetrical. However, any movement will make him fall down and the system (clown + balloon) then loses its symmetry. (B) If the balloon is slightly flat on its base (C–F), then the system is metastable, i.e., a slight perturbation of the clown will not break the symmetry (C, D), whereas a larger perturbation will destabilize the clown (F).Symmetry breaking is ubiquitous in physics, and can lead to phase transitions or pattern formation. It is also an important theme in cell biology, in which polarization is crucial for proper functioning of the cell. Cell polarization typically occurs in response to certain external or internal triggers. A well-known example is chemotaxis, in which a chemical gradient leads to polarization and directed movement of bacterium cells. Polarization also occurs during cytokinesis, in which intracellular stimuli triggered by the mitotic spindle determine the position of the cleavage furrow (Burgess and Chang 2005). Interestingly, cells conserve the ability to polarize even in the absence of an asymmetric signal (Devreotes and Zigmond 1988). For example, chemotactic cells that are presented a uniform concentration of chemoattractant polarize and move in random directions. Another example is blebbing, the spontaneous appearance of bare membrane bulges in some cells.Symmetry breaking in biological systems is a complex phenomenon, because biological systems are always out of equilibrium. Hence, symmetry breaking is not just a transition to a state of lower potential energy. Instead, active, dynamic processes must be considered that feed energy into the system. A biochemical explanation for symmetry breaking was given by Alan Turing. In a seminal paper in 1952 (Turing 1952), he showed that patterns can be generated by simple chemical reactions if the reactants have different diffusion rates. To make this clear, he considered the hypothetical situation in which the morphology of a cell (or cell clump) is determined by two chemical substances (called morphogens). These morphogens also control their own production rate: One enhances morphogen production (the activator) and the other inhibits morphogen production (the inhibitor). It was shown that a spatially homogeneous distribution of morphogens is unstable if the activator diffuses more slowly than the inhibitor. In this case, small stochastic concentration fluctuations are amplified, leading to a chemical instability (“a Turing instability”) and the formation of concentration gradients (or patterns). Reaction–diffusion models of the Turing type have been widely explored to explain polarization and biological development (Gierer and Meinhardt 1972; Sohrmann and Peter 2003; Wedlich-Soldner and Li 2003).Although reaction–diffusion models have proven to be very successful, there is increasing evidence that cell polarization is not only a matter of biochemistry; mechanical aspects play an important role too. Recent work suggests that spontaneous polarization can also be driven by a mechanical instability of the actomyosin cortex of cells. In the remainder of this review, we focus on such mechanical instabilities.     

Wnt Signaling in Bone Development and Disease: Making Stronger Bone with Wnts

The skeleton as an organ is widely distributed throughout the entire vertebrate body. Wnt signaling has emerged to play major roles in almost all aspects of skeletal development and homeostasis. Because abnormal Wnt signaling causes various human skeletal diseases, Wnt signaling has become a focal point of intensive studies in skeletal development and disease. As a result, promising effective therapeutic agents for bone diseases are being developed by targeting the Wnt signaling pathway. Understanding the functional mechanisms of Wnt signaling in skeletal biology and diseases highlights how basic and clinical studies can stimulate each other to push a quick and productive advancement of the entire field. Here we review the current understanding of Wnt signaling in critical aspects of skeletal biology such as bone development, remodeling, mechanotransduction, and fracture healing. We took special efforts to place fundamentally important discoveries in the context of human skeletal diseases.The skeleton has many important functions related to human health. Aside from the classical functions of the skeleton in structural support and movement, the bone matrix forms a major reservoir of calcium and other inorganic ions, and bone cells are active regulators of calcium homeostasis. Recent data suggest that bone cells can secrete hormones (e.g., FGF23 and osteocalcin) and likely play a physiologically significant role in regulating phosphate and energy homeostasis. It has emerged that Wnt signaling plays a major role controlling multiple aspects of skeletal development and maintenance. Thus, understanding how the Wnt pathway controls skeletal growth and homeostasis has broad implications for human health and disease.Cartilage and bone define the skeleton and are produced by chondrocytes and osteoblasts, respectively. During embryonic development, bones are formed by two distinct processes: intramembranous and endochondral ossification (Fig. 1A). A number of cranial bones and the lateral portion of the clavicles are formed by intramembranous ossification. In this process, mesenchymal progenitor cells condense and differentiate directly into bone-forming osteoblasts. The majority of bones in our body are formed by endochondral ossification, during which mesenchymal progenitor cells condense and differentiate first into cartilage-forming chondrocytes to generate an avascular template of the future bone. Chondrocytes in these templates undergo a program of proliferation and progressive cellular maturation. Eventually, they exit the cell cycle and become pre-hypertrophic, then terminally differentiating into hypertrophic chondrocytes, which are eliminated ultimately by apoptosis. Hypertrophic chondrocytes produce a matrix that is calcified and functions as a scaffold for new bone formation. Concomitant with chondrocyte hypertrophy, osteoclasts, osteoblasts, and blood vessels migrate in from perichondral regions and remodel this template into bone.Open in a separate windowFigure 1.Mechanisms of skeleton formation. (A) Bones can form by either intramembranous or endochondral ossification. Both processes are initiated by the condensation of mesenchymal cells. During intramembranous ossification, mesenchymal cells differentiate directly into osteoblasts and deposit bone. During endochondral ossification, mesenchymal cells differentiate into chondrocytes and first make a cartilage intermediate. Chondrocytes in the center of the bone initiate a growth plate, stop proliferating, and undergo hypertrophy. Hypertrophic chondrocytes mineralize their matrix and undergo apoptosis, attracting blood vessels and osteoblasts that remodel the intermediate into bone. (B) The first histologic sign of synovial joint formation is the gathering and flattening of cells, forming the interzone. Cavitation occurs within the presumptive joint separating the two cartilaginous structures. Remodeling and maturation proceed to give rise to the mature synovial joint. Wnt signaling plays a significant role in controlling almost all aspects of skeleton formation. Osteoblasts (purple); chondrocytes (blue); osteochondroprogenitor cells (brown).The developing skeletal elements are often segmented to form joints, which are required to support mobility. Synovial joints, which allow movement via smooth articulation between bones, form when chondrogenic cells in a newly formed cartilage undergo a program of dedifferentiation and flattening to form an interzone (Fig. 1B). Cavitation occurs within the flattened cells, allowing physical separation of the skeletal elements, and the formation of the synovial cavity. Cells and tissues in and around the interzone are remodeled at the same time to form the articular cartilage and other joint structures. Failure to form or maintain joints leads to joint fusion or osteoarthritis, a major skeletal disease.Following its formation, bone remains a regenerative tissue and is maintained during postnatal life by continuous remodeling. This highly active, homeostatic process is required for its functions and is controlled by three cell types: osteoblasts on the bone surface that deposit new bone matrix; osteocytes embedded in bone that are terminally differentiated from osteoblasts and function as mechanical and metabolic sensors; and the matrix-resorbing osteoclasts (Fig. 2). Osteoblasts are derived from mesenchymal stem cells (MSCs), whereas osteoclasts differentiate from hematopoietic progenitors. Decreased bone mass may be due to reduced osteoblast function or elevated osteoclast activity, and, conversely, increased bone mass may result from increased osteoblast function or decreased osteoclast activity. The precise balance of formation and resorption is critical for maintaining normal bone mass, and alterations in this balance lead to common bone diseases such as osteoporosis and osteopetrosis.Open in a separate windowFigure 2.Anatomy of bone. Cortical and trabecular bone represent the two major forms of bone. Osteoblasts (dark purple) are present on the surface and form new bone. Osteocytes (brown) are terminally differentiated osteoblasts that have become embedded in bone and communicate information to one another and to cells on the surface to regulate bone homeostasis. Osteoclasts (blue) are of hematopoietic origin and catabolize bone. A major function of Wnt/β-catenin signaling in osteoblasts is to suppress RANKL and to promote OPG production, thereby inhibiting osteoclast formation.There are two major bone types, cortical and trabecular, which show different anatomical properties (Fig. 2). Cortical (or compact) bone is the solid, densely packed bone that forms the outer layer of most bones and gives strength and rigidity. Trabecular (or cancellous) bone is present mostly in the marrow cavities of long bones and is the dominant bone type in vertebral bodies. Trabecular bone forms a porous, cobweb-like network of trabeculae whose large surface area is thought to facilitate the metabolic activity of bones mediated by osteoblasts and osteoclasts. Trabeculae are sites of active remodeling and will often orient in the direction of mechanical loading, dissipating the energy of loading and adding to bone strength. It is trabecular bone, rather than cortical bone, which is most severely affected in osteoporosis.The Wnt/β-catenin pathway plays a major role in controlling skeletal development and homeostasis, which are the focus of this work. We focus not only on differentiation of skeletal cells and formation of skeletal tissues, but also on the role of the Wnt/β-catenin signaling pathway on bone homeostasis, mechanotransduction, and wound healing, paying particular attention to human and mouse studies.