AUXIN RESPONSE FACTOR17 Is Essential for Pollen Wall Pattern Formation in Arabidopsis

In angiosperms, pollen wall pattern formation is determined by primexine deposition on the microspores. Here, we show that AUXIN RESPONSE FACTOR17 (ARF17) is essential for primexine formation and pollen development in Arabidopsis (Arabidopsis thaliana). The arf17 mutant exhibited a male-sterile phenotype with normal vegetative growth. ARF17 was expressed in microsporocytes and microgametophytes from meiosis to the bicellular microspore stage. Transmission electron microscopy analysis showed that primexine was absent in the arf17 mutant, which leads to pollen wall-patterning defects and pollen degradation. Callose deposition was also significantly reduced in the arf17 mutant, and the expression of CALLOSE SYNTHASE5 (CalS5), the major gene for callose biosynthesis, was approximately 10% that of the wild type. Chromatin immunoprecipitation and electrophoretic mobility shift assays showed that ARF17 can directly bind to the CalS5 promoter. As indicated by the expression of DR5-driven green fluorescent protein, which is an synthetic auxin response reporter, auxin signaling appeared to be specifically impaired in arf17 anthers. Taken together, our results suggest that ARF17 is essential for pollen wall patterning in Arabidopsis by modulating primexine formation at least partially through direct regulation of CalS5 gene expression.In angiosperms, the pollen wall is the most complex plant cell wall. It consists of the inner wall, the intine, and the outer wall, the exine. The exine is further divided into sexine and nexine layers. The sculptured sexine includes three major parts: baculum, tectum, and tryphine (Heslop-Harrison, 1971; Piffanelli et al., 1998; Ariizumi and Toriyama, 2011; Fig. 1A). Production of a functional pollen wall requires the precise spatial and temporal cooperation of gametophytic and sporophytic tissues and metabolic events (Blackmore et al., 2007). The intine layer is controlled gametophytically, while the exine is regulated sporophytically. The sporophytic tapetum cells provide material for pollen wall formation, while primexine determines pollen wall patterning (Heslop-Harrison, 1968).Open in a separate windowFigure 1.Schematic representation of the pollen wall and primexine development. A, The innermost layer adjacent to the plasma membrane is the intine. The bacula (Ba), tectum (Te), and tryphine (T) make up the sexine layer. The nexine is located between the intine and the sexine layers. The exine includes the nexine and sexine layers. B, Primexine (Pr) appears between callose (Cl) and plasma membrane (Pm) at the early tetrad stage (left panel). Subsequently, the plasma membrane becomes undulated (middle panel) and sporopollenin deposits on the peak of the undulated plasma membrane to form bacula and tectum (right panel).After meiosis, four microspores were encased in callose to form a tetrad. Subsequently, the primexine develops between the callose layer and the microspore membrane (Fig. 1B), and the microspore plasma membrane becomes undulated (Fig. 1B; Fitzgerald and Knox, 1995; Southworth and Jernstedt, 1995). Sporopollenin precursors then accumulate on the peak of the undulated microspore membrane to form the bacula and tectum (Fig. 1B; Fitzgerald and Knox, 1995). After callose degradation, individual microspores are released from the tetrad, and the bacula and tectum continue to grow into exine with further sporopollenin deposition (Fitzgerald and Knox, 1995; Blackmore et al., 2007).The callose has been reported to affect primexine deposition and pollen wall pattern formation. The peripheral callose layer, secreted by the microsporocyte, acts as the mold for primexine (Waterkeyn and Bienfait, 1970; Heslop-Harrison, 1971). CALLOSE SYNTHASE5 (CalS5) is the major enzyme responsible for the biosynthesis of the callose peripheral of the tetrad (Dong et al., 2005; Nishikawa et al., 2005). Mutation of Cals5 and abnormal CalS5 pre-mRNA splicing resulted in defective peripheral callose deposition and primexine formation (Dong et al., 2005; Nishikawa et al., 2005; Huang et al., 2013). Besides CalS5, four membrane-associated proteins have also been reported to be involved in primexine formation: DEFECTIVE EXINE FORMATION1 (DEX1; Paxson-Sowders et al., 1997, 2001), NO EXINE FORMATION1 (NEF1; Ariizumi et al., 2004), RUPTURED POLLEN GRAIN1 (RPG1; Guan et al., 2008; Sun et al., 2013), and NO PRIMEXINE AND PLASMA MEMBRANE UNDULATION (NPU; Chang et al., 2012). Mutation of DEX1 results in delayed primexine formation (Paxson-Sowders et al., 2001). The primexine in nef1 is coarse compared with the wild type (Ariizumi et al., 2004). The loss-of-function rpg1 shows reduced primexine deposition (Guan et al., 2008; Sun et al., 2013), while the npu mutant does not deposit any primexine (Chang et al., 2012). Recently, it was reported that Arabidopsis (Arabidopsis thaliana) CYCLIN-DEPENDENT KINASE G1 (CDKG1) associates with the spliceosome to regulate the CalS5 pre-mRNA splicing for pollen wall formation (Huang et al., 2013). Clearly, disrupted primexine deposition leads to aberrant pollen wall patterning and ruptured pollen grains in these mutants.The plant hormone auxin has multiple roles in plant reproductive development (Aloni et al., 2006; Sundberg and Østergaard, 2009). Knocking out the two auxin biosynthesis genes, YUC2 and YUC6, caused an essentially sterile phenotype in Arabidopsis (Cheng et al., 2006). Auxin transport is essential for anther development; defects in auxin flow in anther filaments resulted in abnormal pollen mitosis and pollen development (Feng et al., 2006). Ding et al. (2012) showed that the endoplasmic reticulum-localized auxin transporter PIN8 regulates auxin homeostasis and male gametophyte development in Arabidopsis. Evidence for the localization, biosynthesis, and transport of auxin indicates that auxin regulates anther dehiscence, pollen maturation, and filament elongation during late anther development (Cecchetti et al., 2004, 2008). The role of auxin in pollen wall development has not been reported.The auxin signaling pathway requires the auxin response factor (ARF) family proteins (Quint and Gray, 2006; Guilfoyle and Hagen, 2007; Mockaitis and Estelle, 2008; Vanneste and Friml, 2009). ARF proteins can either activate or repress the expression of target genes by directly binding to auxin response elements (AuxRE; TGTCTC/GAGACA) in the promoters (Ulmasov et al., 1999; Tiwari et al., 2003). The Arabidopsis ARF family contains 23 members. A subgroup in the ARF family, ARF10, ARF16, and ARF17, are targets of miRNA160 (Okushima et al., 2005b; Wang et al., 2005). Plants expressing miR160-resistant ARF17 exhibited pleiotropic developmental defects, including abnormal stamen structure and reduced fertility (Mallory et al., 2005). This indicates a potential role for ARF17 in plant fertility, although the detailed function remains unknown. In addition, ARF17 was also proposed to negatively regulate adventitious root formation (Sorin et al., 2005; Gutierrez et al., 2009), although an ARF17 knockout mutant was not reported and its phenotype is unknown.In this work, we isolated and characterized a loss-of-function mutant of ARF17. Results from cytological observations suggest that ARF17 controls callose biosynthesis and primexine deposition. Consistent with this, the ARF17 protein is highly abundant in microsporocytes and tetrads. Furthermore, we demonstrate that the ARF17 protein is able to bind the promoter region of CalS5. Our results suggest that ARF17 regulates pollen wall pattern formation in Arabidopsis.     

Variability and Constancy in Cellular Growth of Arabidopsis Sepals

Growth of tissues is highly reproducible; yet, growth of individual cells in a tissue is highly variable, and neighboring cells can grow at different rates. We analyzed the growth of epidermal cell lineages in the Arabidopsis (Arabidopsis thaliana) sepal to determine how the growth curves of individual cell lineages relate to one another in a developing tissue. To identify underlying growth trends, we developed a continuous displacement field to predict spatially averaged growth rates. We showed that this displacement field accurately describes the growth of sepal cell lineages and reveals underlying trends within the variability of in vivo cellular growth. We found that the tissue, individual cell lineages, and cell walls all exhibit growth rates that are initially low, accelerate to a maximum, and decrease again. Accordingly, these growth curves can be represented by sigmoid functions. We examined the relationships among the cell lineage growth curves and surprisingly found that all lineages reach the same maximum growth rate relative to their size. However, the cell lineages are not synchronized; each cell lineage reaches this same maximum relative growth rate but at different times. The heterogeneity in observed growth results from shifting the same underlying sigmoid curve in time and scaling by size. Thus, despite the variability in growth observed in our study and others, individual cell lineages in the developing sepal follow similarly shaped growth curves.Cells undergo multiple rounds of growth and division to create reproducible tissues. In some plant tissues, such as expanding cotyledons, reproducibility can occur on a cellular level during specific intervals of development, where cotyledon cells exhibit uniform cellular growth (Zhang et al., 2011). However, several studies on cell division and growth in other developing plant tissues have demonstrated that plant cells exhibit considerable cell-to-cell variability during development (Meyer and Roeder, 2014). For example, in both the Arabidopsis (Arabidopsis thaliana) meristem and leaf epidermis, cells show spatiotemporal variation in individual cell growth rates (GRs; Asl et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012; Uyttewaal et al., 2012). Furthermore, cell divisions have been observed with marked randomness in their timing and orientation (Roeder et al., 2010; Besson and Dumais, 2011; Roeder, 2012). In this study, we identify a hidden, underlying pattern in the seemingly random GR (Box 1) of cells during the formation of sepals in Arabidopsis.Open in a separate windowBox 1.Definitions of GR terms. (For details on the calculations, see “Materials and Methods.”)Plant cell growth is defined as an increase in cell size due to an irreversible expansion of the cell wall. Neighboring cells physically accommodate one another during plant growth because their cell walls are glued together with a pectin-rich middle lamella, which prevents cell mobility. The cell wall is a thin, stiff layer composed of a polymer matrix including cellulose, hemicellulose, and pectin (Somerville et al., 2004; Cosgrove, 2005). Plant cells change their size and shape by modifying their turgor pressure and/or the mechanical properties of their walls, such as elasticity, plasticity, and extensibility. Growing plant cells exert forces on their neighbors through their walls, and cell wall stresses created by these forces feed back to alter the growth anisotropy (Hamant et al., 2008; Sampathkumar et al., 2014). Although these feedbacks can coordinate growth, they may also amplify differences in growth between neighboring cells (Uyttewaal et al., 2012).Two competing computational models have proposed explanations of the cellular heterogeneity observed in growing tissues by making different assumptions about how cells grow. In the first, it is assumed that relative growth rates (RGRs) of all cells are uniform in space and time, whereas variation in the timing of division causes the heterogeneity of cell sizes (Roeder et al., 2010). This model suggests that cell divisions cut the sepal into semiindependent cells, which grow uniformly within the expanding organ (Kaplan and Hagemann, 1991). The second model postulates the reverse process: timing of cell division is uniform, but cellular growth is variable and depends on the size of the cell (Asl et al., 2011). This model suggests that cells are autonomous. Currently, there is biological evidence for both models. Variability in cell division timing is observed in sepals and meristems, whereas variability in cellular GRs has been observed in leaves and meristem cells (Reddy et al., 2004; Roeder et al., 2010; Asl et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012; Uyttewaal et al., 2012). Thus, the debate on how the growth of individual cells within an organ relates to one another remains unresolved.The identification of underlying patterns in noisy cellular growth processes is challenging. Technical difficulties include the capability for cellular-resolution imaging of the tissue at sufficiently small time intervals. Previous studies (Zhang et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012) did not image and track individual cells, or they had a coarse time resolution, with 11- to 48-h intervals between images, which may have hidden important temporal dynamics. We studied growing cells in the Arabidopsis sepal, which allows for live imaging with cellular resolution at 6-h intervals (Roeder et al., 2010). The sepal is the leaf-like outermost floral organ of Arabidopsis (Fig. 1) with four sepals of stereotypical size produced per flower. Its accessibility for live imaging makes the sepal an excellent system for studying organogenesis (Roeder et al., 2010, 2011, 2012; Qu et al., 2014). Sepals exhibit high cellular variability in the timing of division and endoreduplication, an alternative cell cycle in which a cell replicates its DNA but fails to divide (Roeder et al., 2010). Furthermore, quantifying cell growth in sepals may shed light on growth mechanisms of other plant organs, such as leaves (Poethig and Sussex, 1985; Roeder et al., 2010).Open in a separate windowFigure 1.Diverse sizes of Arabidopsis sepal cells. A, Four sepals (s) are the outermost green leaf-like floral organs in Arabidopsis. B and C, Scanning electron micrographs of a mature Arabidopsis sepal show that the outer epidermal cells have a wide range of sizes. Asterisks mark some of the largest cells (giant cells) that can span 1/4 the length of the sepal. Scale = 100 µm.Another key challenge in analyzing cellular growth is the identification of trends in noisy data. Inaccuracies in data acquisition, such as segmentation errors, and noisy growth of individual cells can hide meaningful spatiotemporal trends in growth. GRs measured over longer time intervals will have reduced noise, but they may also obscure important temporal dynamics. Alternatively, previous studies have examined growth of the whole organ or its subregions to avoid cellular noise (De Veylder et al., 2001; Mündermann et al., 2005; Rolland-Lagan et al., 2005, 2014; Kuchen et al., 2012; Remmler and Rolland-Lagan, 2012). However, precise cellular patterns are not resolved. In our study, we use cellular resolution data to define spatially averaged kinematics while keeping the full temporal resolution to identify course-grained spatial trends in the dynamics of cellular growth (Box 1).We analyze the relationships among the growth of individual cell lineages in a developing Arabidopsis sepal by live imaging and computational analyses. We have developed continuous low-order displacement fields to represent the spatially averaged kinematics of the sepal (Box 1). We find that the growth of the tissue surface area, cell lineage area, and wall length follows S curves, suggesting that their GRs vary over time. Additionally, we find that there is a linear correlation between the maximum GR (i.e. size increase per hour) and the size of the cell. We furthermore find that each sepal cell lineage reaches the same maximum RGR (i.e. GR divided by size). However, each cell reaches the maximum RGR at a different time during its development, generating the observed heterogeneity. Thus, we find underlying similarities in the growth curves of sepal cells.     

Herbivore-Induced SABATH Methyltransferases of Maize That Methylate Anthranilic Acid Using S-Adenosyl-l-Methionine

Volatile methyl esters are common constituents of plant volatiles with important functions in plant defense. To study the biosynthesis of these compounds, especially methyl anthranilate and methyl salicylate, we identified a group of methyltransferases that are members of the SABATH enzyme family in maize (Zea mays). In vitro biochemical characterization after bacterial expression revealed three S-adenosyl-l-methionine-dependent methyltransferases with high specificity for anthranilic acid as a substrate. Of these three proteins, Anthranilic Acid Methyltransferase1 (AAMT1) appears to be responsible for most of the S-adenosyl-l-methionine-dependent methyltransferase activity and methyl anthranilate formation observed in maize after herbivore damage. The enzymes may also be involved in the formation of low amounts of methyl salicylate, which are emitted from herbivore-damaged maize. Homology-based structural modeling combined with site-directed mutagenesis identified two amino acid residues, designated tyrosine-246 and glutamine-167 in AAMT1, which are responsible for the high specificity of AAMTs toward anthranilic acid. These residues are conserved in each of the three main clades of the SABATH family, indicating that the carboxyl methyltransferases are functionally separated by these clades. In maize, this gene family has diversified especially toward benzenoid carboxyl methyltransferases that accept anthranilic acid and benzoic acid.Volatile compounds have important roles in the reproduction and defense of plants. Volatiles can attract pollinators and seed dispersers (Dobson and Bergström, 2000; Knudsen et al., 2006) or function as indirect defense compounds that attract natural enemies of herbivores (Dicke, 1994; Degenhardt et al., 2003; Howe and Jander, 2008). A well-studied example for the role of volatiles in plant defense is the tritrophic interaction between maize (Zea mays) plants, their lepidopteran herbivores, and parasitoid wasps of the herbivores. After damage by larvae of Spodoptera species, maize releases a complex volatile blend containing different classes of natural products (Turlings et al., 1990; Turlings and Benrey, 1998a). This volatile blend can be used as a cue by parasitic wasps to find hosts for oviposition (Turlings et al., 1990, 2005). After parasitization, lepidopteran larvae feed less and die upon emergence of the adult wasp, resulting in a considerable reduction in damage to the plant (Hoballah et al., 2002, 2004). The composition of the maize volatile blend is complex, consisting of terpenoids and products of the lipoxygenase pathway, along with three aromatic compounds: indole, methyl anthranilate, and methyl salicylate (Turlings et al., 1990; Degen et al., 2004; Köllner et al., 2004a). In the last decade, several studies have addressed the biosynthesis of terpenoids (Shen et al., 2000; Schnee et al., 2002, 2006; Köllner et al., 2004b, 2008a, 2008b) and indole (Frey et al., 2000, 2004) in maize. The formation of methyl anthranilate and methyl salicylate, however, has not been elucidated.Methyl anthranilate and methyl salicylate are carboxyl methyl esters of anthranilic acid, an intermediate of Trp biosynthesis, and the plant hormone salicylic acid, respectively. Our understanding of methyl anthranilate biosynthesis in plants is very limited. The only enzyme that has been described to be involved in methyl anthranilate synthesis is the anthraniloyl-CoA:methanol acyltransferase in Washington Concord grape (Vitis vinifera; Wang and De Luca, 2005). In contrast, the biosynthesis of methyl salicylate has been well studied in several plant species, such as Clarkia brewerii (Ross et al., 1999), Arabidopsis (Arabidopsis thaliana; Chen et al., 2003), and rice (Oryza sativa; Xu et al., 2006; Koo et al., 2007; Zhao et al., 2010). In all these species, methyl salicylate is synthesized by the action of S-adenosyl-l-methionine:salicylic acid carboxyl methyltransferase (SAMT). The apparent homology of SAMTs from different plant species suggests that methyl salicylate formation in maize, a species closely related to rice, is also catalyzed by an SAMT. SAMT enzymes are considered part of a larger family of methyltransferases called SABATH methyltransferases (D''Auria et al., 2003). The SABATH family also includes methyltransferases producing other methyl esters such as methyl benzoate, methyl jasmonate, and methyl indole-3-acetate (Seo et al., 2001; Effmert et al., 2005; Qin et al., 2005; Song et al., 2005; Zhao et al., 2007). An activity forming methyl anthranilate has not been described in the SABATH family, despite the striking structural similarity between methyl anthranilate and methyl salicylate or methyl benzoate. Two different classes of enzymes, methanol acyl transferases and methyltransferases, therefore, might be responsible for methyl anthranilate biosynthesis in maize (Fig. 1). Some of the SABATH methyltransferases have been shown previously to have methyltransferase activity in vitro using anthranilic acid as substrate (Chen et al., 2003; Zhao et al., 2010), but the biological relevance of such activity is unknown.Open in a separate windowFigure 1.The biosynthesis of methyl anthranilate from anthranilic acid can proceed over two pathways. Pathway A has been documented in grape, while pathway B is demonstrated here. AMAT, Anthraniloyl-CoA:methanol acyltransferase; SAH, S-adenosyl-l-homocysteine.In our ongoing attempt to investigate the biosynthesis and function of maize volatiles, we have studied the biosynthesis of the aromatic methyl esters, methyl salicylate and methyl anthranilate, and their regulation by herbivory. Biochemical characterization of maize benzenoid carboxyl methyltransferases of the SABATH family led to the discovery of a group of anthranilic acid methyltransferases (AAMTs). Homology-based structural modeling combined with site-directed mutagenesis identified the residues critical for the binding of the anthranilic acid substrate. Such functionally important residues are responsible for the diversification and evolution of benzenoid carboxyl methyltransferases in plants.     

Cell biology in neuroscience: Cellular and molecular mechanisms underlying axon formation,growth, and branching

Proper brain wiring during development is pivotal for adult brain function. Neurons display a high degree of polarization both morphologically and functionally, and this polarization requires the segregation of mRNA, proteins, and lipids into the axonal or somatodendritic domains. Recent discoveries have provided insight into many aspects of the cell biology of axonal development including axon specification during neuronal polarization, axon growth, and terminal axon branching during synaptogenesis.

Introduction

Axon development can be divided into three main steps: (1) axon specification during neuronal polarization, (2) axon growth and guidance, and (3) axon branching and presynaptic differentiation (Fig. 1; Barnes and Polleux, 2009; Donahoo and Richards, 2009). These three steps are exemplified during neocortical development in the mouse: upon neurogenesis, newly born neurons engage long-range migration and polarize (Fig. 1, A and B) by adopting a bipolar morphology with a leading and a trailing process (Fig. 1 C). During migration (approximately from embryonic day [E]11 to E18 in the mouse cortex), the trailing process becomes the axon and extends rapidly while being guided to its final destination (lasts until around postnatal day [P]7 in mouse corticofugal axons with distant targets like the spinal cord; Fig. 1, D–F). Finally, upon reaching its target area, extensive axonal branching occurs during the formation of presynaptic contacts with specific postsynaptic partners (during the second and third postnatal week in the mouse cortex; Fig. 1, G–I). Disruption of any of these steps is thought to lead to various neurodevelopmental disorders ranging from mental retardation and infantile epilepsy to autism spectrum disorders (Zoghbi and Bear, 2012). This review will provide an overview of some of the cellular and molecular mechanisms underlying axon specification, growth, and branching.Open in a separate windowFigure 1.Axon specification, growth, and branching during mouse cortical development. Three stages of the development of callosal axons of cortical pyramidal neurons from the superficial layers 2/3 of the somatosensory cortex in the mouse visualized using long-term in utero cortical electroporation. For this class of model axons, development can be divided in three main stages: (1) neurogenesis and axon specification, occurring mostly at embryonic ages (A–C); (2) axon growth/guidance during the first postnatal week (D–F); and (3) axon branching and synapse formation until approximately the end of the third postnatal week (G–I). A, D, and G show coronal sections of mouse cortex at the indicated ages after in utero cortical electroporation of a GFP-coding plasmid at E15.5 in superficial neuron precursors in one brain hemisphere only (GFP signal in inverted color, dotted line indicates the limits of the brain). B, E, and H are a schematic representation of the main morphological changes observed in callosally projecting axons (red) at the corresponding ages. C shows the typical bipolar morphology of a migrating neuron emitting a trailing process (TP) and a leading process (LP) that will ultimately become the axon and dendrite, respectively. F and I show typical axon projections of layer 2/3 neurons located in the primary somatosensory area at P8 and P21, respectively. Neurons and axons in C, F, and I are visualized by GFP expression (inverted color). Image in C is modified from Barnes et al. (2007) with permission from Elsevier. Images in D, F, G, and I are reprinted from Courchet et al. (2013) with permission from Elsevier.

Neuronal polarization and axon specification

Neuronal polarization is the process of breaking symmetry in the newly born cell to create the asymmetry inherent to the formation of the axonal and somatodendritic compartments (Dotti and Banker, 1987). The mechanisms underlying this process have been studied extensively in vitro and more recently in vivo, but the exact sequence of events has remained elusive (Neukirchen and Bradke, 2011) partly because it is studied in various neuronal cell types that might not use the same extrinsic/intrinsic mechanisms to polarize. It is highly likely that at least three factors underlie neuronal polarization: extracellular cues, intracellular signaling cascades, and subcellular organelle localization. The partition-defective proteins (PARs) are a highly conserved family of proteins including two dyads (Par3/Par6 adaptor proteins and the Par4/Par1 serine/threonine kinases) that are required for polarization and axon formation (Shi et al., 2003, 2004; Barnes et al., 2007; Shelly et al., 2007; Chen et al., 2013), while many other intracellular signaling molecules also support axon formation (Oliva et al., 2006; Rašin et al., 2007; Barnes and Polleux, 2009; Shelly et al., 2010; Cheng et al., 2011; Hand and Polleux, 2011; Cheng and Poo, 2012; Gärtner et al., 2012). These intracellular signaling pathways are influenced by localized extracellular cues that instruct which neurite becomes the axon by either directly promoting axon extension or repressing axon growth in favor of dendritic growth (Adler et al., 2006; Yi et al., 2010; Randlett et al., 2011b; Shelly et al., 2011).The role of organelle subcellular localization during neuronal polarization is a more controversial issue. Initially, the orientation of organelles, including the Golgi complex, centrosomes, mitochondria, and endosomes, was shown to correlate with the neurite that becomes the axon in vitro (Bradke and Dotti, 1997; de Anda et al., 2005, 2010) and in vivo (de Anda et al., 2010). However, more recent studies suggest that the positioning of the centrosome is not necessary for neuronal polarization (Distel et al., 2010; Nguyen et al., 2011). Centrosome localization is likely constrained by microtubule organization within the cell, and therefore the centrosome position within the cell changes dynamically during different stages of polarization (Sakakibara et al., 2013). The question of how the interplay between extracellular cues, intracellular signaling, and organelle localization lead to polarization has pushed the field to perform more extensive in vivo imaging studies as in vitro systems/models have a difficult time recapitulating the complex environment and rely on neurons that were previously polarized in vivo.Like other epithelial cells, neural progenitors present a high degree of polarization along the apico-basal axis (Götz and Huttner, 2005). One of the major questions still needing to be addressed is how, or if, newly born mammalian neurons inherit some level of asymmetry from their parent progenitors (Barnes and Polleux, 2009). Recent studies have attempted to answer this question in vivo but have found that the answer might vary in each neuronal subtype. Retinal ganglion cells (RGCs), retinal bipolar neurons, and tegmental hindbrain nuclei neurons seem to inherit the apical/basolateral polarity from their progenitors (Morgan et al., 2006; Zolessi et al., 2006; Distel et al., 2010; Randlett et al., 2011a). In cortical neurons, hippocampal neurons, and cerebellar granule neurons, this relationship is unclear, in part because newly born cortical neurons first exhibit a multipolar morphology with dynamic neurites emerging from the cell body before adopting a bipolar morphology, suggesting they may not retain a predisposed parental polarity (Hand et al., 2005; Barnes et al., 2007). Other factors also suggest that different neuronal subtypes use different mechanisms during polarization. One such factor is the position where neurons specify their axon relative to the original apical/basolateral axis of their progenitors. As an example, cortical neurons in the mouse brain protrude an axon from the membrane facing the original apical surface toward the ventricular zone (Hand et al., 2005; Barnes et al., 2007; Shelly et al., 2007), whereas zebrafish RGCs form their axon from the membrane on the basolateral side (Zolessi et al., 2006; Randlett et al., 2011b). Another significant difference between cortical neurons and RGCs is related to the timing of axogenesis and dendrogenesis. RGCs tend to form their axons and dendrites at the same time during migration (Zolessi et al., 2006; Randlett et al., 2011b). However, cortical neurons form a long axon during migration before significant dendrite arborization takes place. These differences in the regulation of polarization and sequence of axon versus dendrite outgrowth may be linked to the localization of extracellular cues relative to the immature neuron during polarization (Yi et al., 2010).

Neuronal polarization, cytoskeletal dynamics, and polarized transport

What exactly makes the axonal compartment distinct from the somatodendritic domain? This can most easily be illustrated by focusing on the cytoskeleton that forms the framework of the developing axon. The cytoskeleton is composed of microtubules, actin filaments, and intermediate filaments (also called neurofilaments) along with their associated binding partners. Microtubules are composed of α- and β-tubulin subunits that polymerize to form a long filament intrinsically polarized by the addition of tubulin subunits to only one side of the growing filament called the plus end, while on the opposite side depolymerization occurs. It was discovered more than thirty years ago that the axon of a neuron contains a very uniform distribution of microtubules with the plus end facing away from the cell body (Heidemann et al., 1981). Through the years this observation was confirmed in many neuron cell types, and it was determined that dendrites do not have this uniform plus-end out network of microtubules (Fig. 2; Baas et al., 1988). Dendrites appear to have a complex array of microtubule orientations that may vary between species and/or neuronal subtypes. Current research shows that proximal dendrites are composed of mainly minus-end out microtubules, whereas more distal dendrites transition from an equal distribution of minus-end out and plus-end out microtubules to mainly plus-end out microtubules (Stone et al., 2008; Yin et al., 2011; Ori-McKenney et al., 2012). The orientation of microtubules matters greatly because it determines the relative contribution of microtubule-dependent motor proteins (kinesins and dyneins), which are the main motor proteins carrying various cargoes within cells and in particular are responsible for long-range transport in very large cells such as neurons. Dynein (a minus end–directed microtubule motor) is known to be responsible for both the transport of microtubules away from the cell body and for the uniform polarity of microtubules in the axon (Ahmad et al., 1998; Zheng et al., 2008). Recently, it was discovered that kinesin-1 (a plus end–directed microtubule motor) is required for the minus-end out orientation of microtubules in the dendrites of Caenorhabditis elegans DA9 neurons through selective transport of plus-end out microtubule fragments out of the dendrite (Yan et al., 2013). Another hallmark that differentiates the axonal and somatodendritic compartments is the microtubule-associated proteins (MAPs) that decorate microtubules to regulate their bundling and stability (Hirokawa et al., 2010). Microtubules in the axon are mainly decorated by Tau and MAP1B, whereas microtubules in the dendrites are labeled by proteins of the MAP2a-c family. The role of Tau in axon elongation remains controversial because early reports (Harada et al., 1994; Tint et al., 1998; Dawson et al., 2001) of Tau knockout alone suggested that axons were unaffected, but this apparent lack of phenotype might originate from the functional redundancy between MAPs as Tau/MAP1b double knockout mice show clear axon growth defects (Takei et al., 2000).Open in a separate windowFigure 2.Polarity maintenance and trafficking of somatodendritic and axonal proteins. Neurons are polarized into two main compartments: the somatodendritic domain and the axon. These domains are characterized by the underlying cytoskeleton and their unique protein signatures. The axonal cytoskeleton is defined by its uniform microtubule orientation where each microtubule is oriented with its plus end away from the cell body, while the dendrites contain a mixture of microtubules oriented in both directions. The proximal axon is characterized by a structure known as the axon initial segment (AIS, see inset). This highly ordered structure creates a diffusion barrier between the axonal compartment and the rest of the cell. F-actin is responsible for the cytoplasmic barrier, while sodium channels anchored by Ankyrin G form the basis of the plasma membrane barrier. Tau is retained in the axon by a microtubule-based filter at the AIS. Molecular motors (including kinesin, dynein, and myosin) then use the underlying cytoskeleton to restrict cargo transport to either the axon (such as Cntn1 and L1) or the dendrites (such as PSD95, AMPARs, and NMDARs).The dynamics of actin polymerization into actin filaments (F-actin) also play an important role in defining the axonal compartment, and contain an intrinsic polarity based on the polymerization of the free G-actin subunits (Hirokawa et al., 2010). Beyond the well-documented early role of F-actin dynamics in neurite outgrowth, multiple groups have shown that the disruption of actin polymerization allows dendritically localized proteins to incorrectly enter the axonal compartment (Winckler et al., 1999; Lewis et al., 2009; Song et al., 2009). The existence of a “diffusion barrier” in the proximal part of newly formed axons (Song et al., 2009) was long suspected. One of the current hypotheses is that a dense F-actin meshwork creates a cytoplasmic diffusion barrier shortly after polarization, which in part separates the axonal compartment from the neuronal cell body (Fig. 2, inset). Based on functional analysis and electron microscopy analysis, this “F-actin–based filter” is oriented so that the plus ends point toward the cell body while the minus ends point into the axon (Lewis et al., 2009, 2011; Watanabe et al., 2012). Two recent papers show via high resolution imaging techniques that indeed the axon has a unique F-actin network that is not found in dendrites (Watanabe et al., 2012; Xu et al., 2013). The development of this F-actin meshwork appears to directly precede the formation of the axon initial segment (AIS; Song et al., 2009; Galiano et al., 2012). An intra-axonal diffusion barrier, composed of Spectrins and Ankyrin B, defines the eventual position of the AIS. This boundary excludes Ankyrin G, which instead clusters in the most proximal part of the axon close to the cell body, where the AIS will form (Galiano et al., 2012). Ankyrin G is required for AIS formation and maintenance, and its loss causes the axon to start forming protrusions resembling dendritic spines (Hedstrom et al., 2008). Microtubules also play an important role at the AIS, as recent evidence suggests that Tau is retained in the axon through a microtubule-based diffusion barrier independently of the F-actin based filter (X. Li et al., 2011). The AIS is important in the formation of a plasma membrane barrier between the axonal and somatodendritic domains and its disruption affects both neuronal polarity and function because it is critical for clustering of voltage-dependent sodium channels and action potential initiation (Rasband, 2010).One of the critical cellular mechanisms underlying neuronal polarization is the polarized transport of various cargoes in axons and dendrites. Transport of proteins and various organelles is performed by the microtubule-dependent motor proteins kinesin and dynein (Hirokawa et al., 2010). Studies from many laboratories have demonstrated that kinesin motors can carry cargo to both the axonal and dendritic compartments (Burack et al., 2000; Nakata and Hirokawa, 2003). The mechanism for how selection occurs is not completely understood, but it probably incorporates both the affinity of the kinesin head for microtubules and the cargo bound to the motor protein (Nakata and Hirokawa, 2003; Song et al., 2009; Jenkins et al., 2012). In the axon, dynein works to bring cargo and retrograde signals back to the cell body, whereas in the dendrites it is responsible for much of the transport from the soma into the dendrites (Zheng et al., 2008; Kapitein et al., 2010; Harrington and Ginty, 2013). Additionally, the F-actin–dependent myosin motors can affect the polarized transport of cargos by using the F-actin–based cytoplasmic filter at the AIS to deny or facilitate entry of vesicles into the axon. Loss of the actin filter or myosin Va activity (a plus end–directed motor) allows dendritic cargos into the axon, whereas myosin VI (a minus end–directed motor) both removes axonal proteins from the dendritic surface and funnels vesicles containing axonal proteins through the actin filter at the AIS (Lewis et al., 2009, 2011; Al-Bassam et al., 2012). A current working hypothesis is that vesicles composed of multiple cargoes contain binding sites for each of these motors, and that through unknown mechanisms the activity of the motors can be differentially regulated to control the directionality of transport. An interesting example of how the interplay between different motors and cargo adaptors could lead to polarized transport was recently described for mitochondria (van Spronsen et al., 2013).

Axon growth

Microtubule dynamics regulate axon growth.

After axon specification, axon growth constitutes the second step of axonal development and is tightly linked to axon guidance toward the proper postsynaptic targets. Axon elongation by the growth cone is the product of two opposite forces (Fig. 3): slow axonal transport and the polymerization of microtubules providing a pushing force from the axon shaft, and the retrograde flow of actin providing a pulling force at the front of the growth cone (Letourneau et al., 1987; Suter and Miller, 2011). Although coordinated actin and microtubule dynamics are required for the proper function of the growth cone, it was reported that agents disrupting the actin cytoskeleton have limited consequences on axon elongation and are rather involved in axon guidance in vitro (Marsh and Letourneau, 1984; Ruthel and Hollenbeck, 2000) and in vivo (Bentley and Toroian-Raymond, 1986). Furthermore, local disruption of actin organization in the growth cone of minor neurites allows them to turn into axons (Bradke and Dotti, 1999; Kunda et al., 2001), indicating that the dense actin network present at the periphery of an immature neuronal cell body and in immature neurites may prevent microtubule protrusion and elongation necessary for axon specification.Open in a separate windowFigure 3.Cytoskeletal changes during axon elongation and branching. Representation of axon elongation and collateral branch formation in a cultured neuron. Axon growth is a discontinuous process, and collateral branches often originate from sites where the growth cone paused (gray dotted line), after it has resumed its progression. Other modalities of branch formation can occur through the formation of filopodia and lamellipodia. Red box shows a magnification of the main growth cone. Microtubules from the axon shaft spread into the central (C) zone. Some microtubules pass through the transition (T) zone, containing F-actin arcs, to explore filopodia from the peripheral (P) zone. Upon the proper stimulation by extracellular guidance cues or growth-promoting cues, microtubules are stabilized and invade the P-zone where they provide a pushing force, which, combined with the traction force from the actin treadmilling, provides the force required for growth cone extension. Green box shows the cytoskeletal changes occurring during collateral branch formation in the axon. Filopodia and lamellipodia are primarily F-actin–based protrusions that get invaded by microtubules, then elongate upon microtubule bundling. At later developmental stages, axon branches are stabilized or retracted (blue box) by mechanisms relying on the access to extracellular neurotrophins and/or neuronal activity and synapse formation.Contrary to actin, microtubule polymerization is required to sustain axon elongation and branching (Letourneau et al., 1987; Baas and Ahmad, 1993). Axonal proteins and cytoskeletal elements are transported along the axon through slow axonal transport by molecular motors (Yabe et al., 1999; Xia et al., 2003). It is still controversial whether tubulin and other cytoskeletal elements are transported in the axon as monomers and/or as polymers (Roy et al., 2000; Terada et al., 2000; Wang et al., 2000; Brown, 2003; Terada, 2003). Nonetheless, disruption of the slow transport of tubulin impairs the pushing force resulting from microtubule polymerization and impairs axon elongation (Suter and Miller, 2011). Therefore, it is not surprising that axon growth is affected in vitro and in vivo by disruption of plus-end microtubule-binding proteins such as APC (Shi et al., 2004; Zhou et al., 2004; Yokota et al., 2009; Chen et al., 2011) or EB1 and EB3 (Zhou et al., 2004; Jiménez-Mateos et al., 2005; Geraldo et al., 2008), microtubule-associated proteins such as MAP1B (Black et al., 1994; Takei et al., 2000; Dajas-Bailador et al., 2012; Tortosa et al., 2013), or proteins regulating microtubule severing and reorganization such as KIF2A (Homma et al., 2003), katanin, and spastin (Karabay et al., 2004; Yu et al., 2005; Wood et al., 2006; Butler et al., 2010).The contribution of microtubule dynamics to axon growth is not limited to growth cone dynamics but also involves axonal transport and polymerization along the axon shaft. Moreover, changing the balance between microtubule stabilization and depolymerization by local application of microtubule stabilizing agents is sufficient to instruct one neurite to grow and adopt an axon fate (Witte et al., 2008). Many kinase pathways converge on Tau and other axonal MAPs to regulate their function by phosphorylation (Morris et al., 2011). Among them, the MARK kinases regulate microtubule stability and axonal transport through phosphorylation of Tau (Drewes et al., 1997; Mandelkow et al., 2004). Interestingly, MARK-related kinases SAD-A/B control axon specification in part through phosphorylation of Tau (Barnes et al., 2007) and have very recently been linked to the growth and branching of the axons of sensory neurons (Lilley et al., 2013). Our work recently demonstrated that another family member related to MARKs and SAD kinases, called NUAK1, controls axon branching of mouse cortical neurons through the regulation of presynaptic mitochondria capture (Courchet et al., 2013). To what extent the regulation of Tau and other MAPs by the MARKs, SADs, and NUAK1 kinases contributes to axon elongation remains to be explored.

Where does axon elongation take place?

Growth cone progression and guidance constitute the main driver of axonal growth during development, but this process is unlikely to account for the totality of axon elongation. This is especially true after the axon has reached its final target but the axon shaft keeps growing in proportion to the rest of the body. One mechanism that may contribute to this “interstitial” form of axon elongation during brain/body size increase (see Fig. 1 for an example during postnatal cortex growth) is axon stretching, a process that can induce axon elongation in vitro (Smith et al., 2001; Pfister et al., 2004; Loverde et al., 2011) and in vivo (Abe et al., 2004). Aside from extreme stretching performed through the application of external forces, stretching could also contribute to the natural elongation of the axon in response to the tension resulting from growth cone progression (Suter and Miller, 2011).Axon elongation requires the addition of new lipids, proteins, cytoskeleton elements, and organelles along the axon. Where does the synthesis and incorporation of new components take place? Polysaccharides and cholesterol synthesis mostly occur in the cell body; however, lipid synthesis and/or incorporation can occur along the axon as well (Posse De Chaves et al., 2000; Hayashi et al., 2004). The growth cone is also a site of endocytosis, membrane recycling, and exocytosis (Kamiguchi and Yoshihara, 2001; Winckler and Yap, 2011; Nakazawa et al., 2012). One of the best studied examples of endocytosis and its role in axon growth and neuronal survival is the retrograde trafficking of TrkA receptor by target-derived nerve growth factor (NGF) in the peripheral nervous system (Harrington and Ginty, 2013).

Axon branching and presynaptic differentiation

Where do axon branches form?

The last step of axon development is terminal branching, which allows a single axon to connect to a broad set of postsynaptic targets. Collateral branches are formed along the axon through two distinct mechanisms: the first modality of branching is through splitting or bifurcation of the growth cone, which is linked to axon guidance and to the capacity of one single neuron to reach two targets that are far apart with one single axon. Growth cone splitting is observed in vivo in various neuron types including cortical neurons (Sato et al., 1994; Bastmeyer and O’Leary, 1996; Dent et al., 1999; Tang and Kalil, 2005), sympathetic neurons (Letourneau et al., 1986), motorneurons (Matheson and Levine, 1999), sensory neurons (Ma and Tessier-Lavigne, 2007), or mushroom body neurons in Drosophila (Wang et al., 2002). The second modality, known as interstitial branching, occurs through the formation of collateral branches directly along the axon shaft. Contrary to growth cone splitting, interstitial branching serves the purpose of raising axon coverage locally in order to define their “presynaptic territory”, and may contribute to increased network connectivity (Portera-Cailliau et al., 2005). Although both mechanisms can occur simultaneously in the same neuron, the relative importance of splitting versus interstitial branching is highly divergent from one neuron type to the other (Bastmeyer and O’Leary, 1996; Matheson and Levine, 1999; Portera-Cailliau et al., 2005).In culture, the axon grows in a non-continuous fashion with frequent growth cone pausing. Time-lapse imaging of sensorimotor neurons revealed that interstitial branching often occurs at the site where the growth cone paused, shortly after it has continued its course (Szebenyi et al., 1998). Accordingly, treatments with neurotrophins that slow the growth cone correlate with increased axon branching (Szebenyi et al., 1998). This suggests that growth cone pausing leaves a “mark” along the axon shaft that might predetermine future sites of branching (Kalil et al., 2000). However, local applications of neurotrophins shows that aside from growth cone pause sites the axon shaft remains competent to form collateral branches upon stimulation by extracellular factors (Gallo and Letourneau, 1998; Szebenyi et al., 2001), through the formation of filopodia or lamellipodia. Similar observations in vivo revealed that cortical axons are highly dynamic during development and form multiple filopodia that are the origin of collateral branches (Bastmeyer and O’Leary, 1996). Lamellipodia can be observed as motile, F-actin–dependent “waves” along the axon in vitro (Ruthel and Banker, 1998) and in vivo (Flynn et al., 2009). Moreover, disruption of microtubule organization impairs lamellipodia formation along the axon and is correlated with decreased axon branching (Dent and Kalil, 2001; Tint et al., 2009).

Cytoskeleton dynamics and axon branch formation.

Regardless of what type of protrusion gives rise to a branch, cytoskeletal reorganization in the nascent branch generally follows a similar sequence (Fig. 3): initially F-actin filament reorganization gives rise to a protrusion (filopodia, lamellipodia), followed by microtubule invasion of this otherwise transient structure to consolidate it, before the mature branch starts elongating through microtubule bundling (Gallo, 2011). Actin filaments accumulate along the axon and form “patches” that serve as nucleators for axon protrusions such as filopodia and lamellipodia (Korobova and Svitkina, 2008; Mingorance-Le Meur and O’Connor, 2009; Ketschek and Gallo, 2010). The mRNA for β-actin and regulators of actin polymerization such as Wave1 or Cortactin accumulate along the axons of sensory neurons and form hot-spots of local translation that are associated to NGF-dependent branching (Spillane et al., 2012; Donnelly et al., 2013). Subsequently, microtubules in the axon shaft undergo fragmentation at branch points as a prelude to branch invasion by microtubules (Yu et al., 1994, 2008; Gallo and Letourneau, 1998; Dent et al., 1999; Hu et al., 2012), a process that may disrupt transport locally to help trap molecules and organelles at branch points. Moreover, severed microtubules are transported into branches, a process required for branch stabilization (Gallo and Letourneau, 1999; Ahmad et al., 2006; Qiang et al., 2010; Hu et al., 2012). Interestingly, it is clear that, just like growth cone–mediated axon elongation, F-actin and microtubule reorganization events are interconnected to sustain axon branching (Dent and Kalil, 2001). As an example, microtubule-severing enzymes can also contribute to actin nucleation and filopodia formation (Hu et al., 2012).

Is axon branching linked to axon elongation?

Like in the growth cone, cytoskeleton reorganization constitutes the backbone of branch formation. It is therefore not surprising that many manipulations of the cytoskeleton affect both axon elongation and branch formation (Homma et al., 2003; Chen et al., 2011). Moreover, conditions that primarily disrupt axon elongation could secondarily disrupt branching by impairing the ability of the nascent branch to grow. However, axon elongation and axon branching can be considered as two separate phenomena and can be operationally separated because conditions disrupting one do not systematically affect the other. As an example, the microtubule-severing proteins katanin and spastin have differential consequences on axon elongation (primarily dependent upon katanin function) and branching (mostly spastin mediated; Qiang et al., 2010), taxol treatment (which stabilizes microtubules) affects axon elongation but not branching (Gallo and Letourneau, 1999), and disruption of TrkA endocytosis by knock-down of Unc51-like kinase (ULK1/2) proteins has opposite effects on axon elongation and branching (Zhou et al., 2007). In vivo, superficial layer cortical neurons initially go through a phase of elongation through the corpus callosum without branching (see Fig. 1), then stop elongating and form collateral branches in the contralateral cortex (Mizuno et al., 2007; Wang et al., 2007). It is conceivable that even before myelination, axons are actively prevented from branching at places and stages when they elongate (for example in the white matter of the neocortex) where they tend to be highly fasciculated. The identities of the molecules that inhibit interstitial branching along the axon shaft are currently unknown.

Regulation of axon branching by activity.

Immature neurons display spontaneous activity in the form of calcium waves (Gu et al., 1994; Gomez and Spitzer, 1999; Gomez et al., 2001) and spontaneous vesicular release long before they have completed axon development, which suggested a role for early neuronal activity in axon development and guidance (Catalano and Shatz, 1998). Cell-autonomous silencing of neurons in vivo by transfection of the hyperpolarizing inward-rectifying potassium channel Kir2.1 in olfactory neurons (Yu et al., 2004), in RGCs (Hua et al., 2005) or in cortical pyramidal neurons (Mizuno et al., 2007; Wang et al., 2007), or in vitro through infusion of tetrodotoxin (which blocks action potentials generation) in co-cultures of thalamo-cortical projecting neurons (Uesaka et al., 2007) results in a decrease in terminal axon branching, indicating that synaptic activity is required for axons to fully develop their branching pattern. Moreover, inhibition of synaptic release by expression of tetanus toxin light chain (TeTN-LC; Wang et al., 2007) also abolished terminal axon branching, suggesting that the formation of functional presynaptic release sites is required cell autonomously to control terminal axon branching. However, one potential limitation of the experiments involving TeTN-LC is that it blocks most VAMP-mediated vesicular release (with the exception of VAMP7, also called tetanus toxin–independent VAMP, or TI-VAMP). Therefore TeTN-LC action may not be limited to blocking synaptic vesicle release, but could also inhibit peptide release through vesicles containing neurotrophins for example, or other important trophic factors required for axon branching. More recent experiments through silencing of postsynaptic targets revealed that branching of callosal or thalamocortical axons is also dependent upon the activity of the postsynaptic targets (Mizuno et al., 2010; Yamada et al., 2010), albeit activity of the presynaptic neuron is required earlier during the branching process than activity of the postsynaptic targets (Mizuno et al., 2010). Activity is also required in some neurons at the phase of axon elongation through a feedback loop involving the activity-dependent up-regulation of guidance molecules (Mire et al., 2012).How much does spontaneous or evoked neuronal activity contribute to branching? Reduction of neuronal activity through hyperpolarization induced by overexpression of Kir2.1 significantly reduces axon branching without completely eliminating it (Hua et al., 2005; Mizuno et al., 2007; Wang et al., 2007). Activity seems to serve as a competitive regulator of axon branching with regard to its neighbors because silencing of neighboring axons restores normal branching (Hua et al., 2005). Interestingly, neuronal activity induces neurotrophin expression locally, suggesting that activity can contribute to branching partly through activation of activity-independent branching mechanisms (Calinescu et al., 2011).Neuronal activity can regulate branching through modification of the actin cytoskeleton via RhoA activation (Ohnami et al., 2008), and mRNA accumulates at presynaptic sites, indicating a correlation between local translation and synaptic activity (Lyles et al., 2006; Taylor et al., 2013). Neuronal activity is associated with changes in intracellular Ca²⁺ signaling, which has been shown to play a deterministic function in axon growth (Gomez and Spitzer, 1999). Calcium signaling activates the Ca²⁺/calmodulin-dependent kinases (CAMKs) that are known to regulate axon branching in vitro (Wayman et al., 2004; Ageta-Ishihara et al., 2009) and in vivo (Ageta-Ishihara et al., 2009).

Stabilization and refinement of the axonal arborization.

Axon branches are often formed in excess during development, then later refined to select for specific neural circuits (Luo and O’Leary, 2005). Long-range axon branch retraction has long been observed in layer V cortical neurons that initially project to the midbrain, hindbrain, and spinal cord (O’Leary and Terashima, 1988; Bastmeyer and O’Leary, 1996). At later stages, pyramidal neurons from the primary visual cortex will retract their spinal projection through axon pruning, whereas pyramidal neurons from the primary motor cortex will stabilize this projection but retract their axonal branches growing toward visual targets such as the superior colliculus. The molecular mechanisms controlling this area-specific pattern of axon branch pruning are still poorly understood, but seem to involve extracellular cues such as semaphorins and Rac1-dependent signaling (Bagri et al., 2003; Low et al., 2008; Riccomagno et al., 2012). Another example is the well-characterized refinement of retino-geniculate axons during the selective elimination of binocular input of RGC axon synapses onto relay neurons in the dorsal lateral geniculate nucleus (Muir-Robinson et al., 2002). Interestingly, some axons use caspase-dependent pathways locally to induce the selective retraction of axon branches during the process of pruning (Nikolaev et al., 2009; Simon et al., 2012).Circuit refinement and selective branch retraction can be observed in vivo at the level of the neuromuscular junction where individual branches of motor axons are eliminated asynchronously (Keller-Peck et al., 2001). In the developing CNS, neurotrophin-induced branch retraction can also be observed in a context of competition between neighboring axons (Singh et al., 2008). One other way of stabilizing axon branches is through the formation of synapses with postsynaptic targets. In the visual system, the initial axon arbor is refined to establish ocular dominance through activity-dependent retraction of less active branches (Ruthazer et al., 2003). Time-lapse imaging of RGC axons in zebrafish or in Xenopus tadpole revealed that the formation of presynaptic sites occurs concomitantly to axon branching, and branches that form presynaptic structures are less likely to retract (Meyer and Smith, 2006; Ruthazer et al., 2006). The stabilization of axon branches through formation of synaptic contacts parallels with the stabilization of dendritic branches through synapse formation and stabilization (Niell et al., 2004; J. Li et al., 2011). The role of presynaptic bouton formation goes beyond the stabilization of axonal branches because in vivo, new axon branches can emerge from existing presynaptic terminals (Alsina et al., 2001; Javaherian and Cline, 2005; Panzer et al., 2006).In conclusion, axon growth and branching can be regulated by both activity-dependent and activity-independent mechanisms during development. However, for mammalian CNS axons, much more work is needed to define (1) the precise molecular mechanisms underlying axon branching; (2) the cellular and molecular mechanisms regulating the key transition between axon growth and branching when axons start forming presynaptic contacts with their postsynaptic partners; and (3) the mechanisms regulating axon pruning during synapse elimination.     

Rab GTPases and membrane identity: Causal or inconsequential?

Rab GTPases are highly conserved components of vesicle trafficking pathways that help to ensure the fusion of a vesicle with a specific target organelle membrane. Specific regulatory pathways promote kinetic proofreading of membrane surfaces by Rab GTPases, and permit accumulation of active Rabs only at the required sites. Emerging evidence indicates that Rab activation and inactivation are under complex feedback control, suggesting that ultrasensitivity and bistability, principles established for other cellular regulatory networks, may also apply to Rab regulation. Such systems can promote the rapid membrane accumulation and removal of Rabs to create time-limited membrane domains with a unique composition, and can explain how Rabs define the identity of vesicle and organelle membranes.

Rab GTPases regulate membrane tethering and vesicle fusion

Eukaryotic cells are defined in part by their complex membrane organelles. This organization permits the coexistence of different chemical environments within the same cell. For example, the endoplasmic reticulum (ER) is a neutral pH, reducing environment containing chaperones conducive to protein folding and the formation of disulfide bonds, whereas the lysosomes are ∼pH 5 and contain catabolic enzymes maximally active at acidic pH. Though valuable, this organization requires some form of active transport machinery for the exchange of material between these compartments because large hydrophilic molecules such as proteins cannot easily cross membranes. This transfer of molecules between compartments is achieved by vesicular transport systems that use cytosolic coat protein complexes to select small regions of membrane and shape these into defined 40–80-nm-diameter transport vesicles (Bonifacino and Glick, 2004; Faini et al., 2013). Vesicle coats contain binding sites for specific transport sequences, and thus only transfer a subset of proteins into the vesicle. Once produced, these vesicles have to identify, tether to, and then fuse with a specific target organelle (Zerial and McBride, 2001). Research over many years has defined small transmembrane proteins (SNAREs) and a set of accessory factors as the minimal machinery for membrane fusion (McNew et al., 2000; Shi et al., 2012). Tethering is a less well-defined event involving the Rab GTPases and effector protein complexes, typically large extended molecules thought to bridge the space between two approaching membranes (Gillingham and Munro, 2003).Rab GTPases were first linked to vesicle transport by groundbreaking genetic screens for mutants defective in protein secretion (Novick et al., 1980; Salminen and Novick, 1987). Sec4, Rab8 in humans, was found to function in the terminal step of the secretory pathway, delivery of Golgi-derived transport vesicles to the cell surface (Salminen and Novick, 1987; Goud et al., 1988). Ypt1, Rab1 in humans, was then shown to regulate secretion at the Golgi apparatus (Segev et al., 1988; Bacon et al., 1989). These findings led to an influential model for Rab function in which the cycle of GTPase activation and inactivation is coupled to recognition events in vesicle docking (Bourne, 1988). Consistent with the idea that they control vesicle targeting, work in mammalian cells then showed that there is a large family of highly conserved Rab GTPases, each with a specific subcellular localization (Chavrier et al., 1990). A series of seminal studies has since provided direct evidence that Rab1 and Rab5 promote membrane fusion (Gorvel et al., 1991; Segev, 1991) by regulating the activation and engagement of SNAREs (Lian et al., 1994; Søgaard et al., 1994), as a consequence of recruiting tethering factors to membrane surfaces (Segev, 1991; Sapperstein et al., 1996; Cao et al., 1998; Christoforidis et al., 1999; McBride et al., 1999; Allan et al., 2000; Shorter et al., 2002). Similar findings were also made for the Rab Ypt7, which functions in vacuole docking in yeast (Price et al., 2000; Ungermann et al., 2000), a system that allows direct visualization of docked or tethered intermediates due to the large size of the membrane structures (Wang et al., 2002).The evidence that Rabs function upstream of SNARE protein in vesicle trafficking pathways has led to the notion that Rabs help to define the identity of vesicle and organelle membranes (Pfeffer, 2001; Zerial and McBride, 2001). This is best exemplified by the early endocytic pathway, where the identity of early and late endosomes is thought to be determined by Rab5 and Rab7, respectively (Rink et al., 2005). However, in most other cases it remains unclear if this is a causal relationship, where the Rab directly defines the identity of the membrane rather than acting as an upstream regulator of vesicle targeting before the SNARE-mediated membrane fusion event. In addition to Rabs, GTPases of the Arf/Arl family and specific phosphoinositide lipids have also been proposed to act in specifying membrane identity (Munro, 2002; Di Paolo and De Camilli, 2006). It therefore seems likely that no single factor can explain how membrane identity is achieved in vesicle transport, and that Rabs, phosphoinositides, and other factors act in concert.

Rab GEFs provide the minimal machinery for targeting and activation

Despite the progress in defining Rab function, the claim that Rab GTPases define organelle identity therefore remains premature due to crucial unanswered questions. In particular, the issue of how Rabs are targeted to specific organelles, or even restricted to subdomains of these organelles, has remained problematic. Initial work using chimeric GTPases suggested that the variable C-terminal region of the different Rabs provided a targeting mechanism (Chavrier et al., 1991). However, subsequent work indicated that this failed to provide a general mechanism to explain specific Rab targeting, and that multiple regions of the Rab including C-terminal prenylation contribute to membrane recruitment (Ali et al., 2004). Emerging evidence based on the improved understanding of the family of Rab guanine nucleotide exchange factors (GEFs) now provides an alternative view for Rab activation at specific membrane surfaces. Mechanistic details of how Rab GEFs activate Rabs have been discussed elsewhere (Barr and Lambright, 2010), and are not directly relevant for this discussion so won’t be detailed further. Two studies now show that Rab GEFs can provide the minimal machinery needed to target a Rab to a specific membrane within the cell (Gerondopoulos et al., 2012; Blümer et al., 2013). In both cases, Rab GEFs were fused to mitochondrial outer membrane targeting sequences, and the effects on different Rabs observed. Using this strategy it was possible to specifically target Rab1, Rab5, Rab8, Rab35, and Rab32/38 to mitochondria with biochemically defined cognate GEFs (Gerondopoulos et al., 2012; Blümer et al., 2013). Mutants that either reduced the nucleotide exchange activity of the GEF or the target GTPase gave a correspondingly reduced rate of Rab targeting (Blümer et al., 2013). Alone this does not provide a full explanation for Rab targeting; for this an understanding of the interaction of prenylated Rabs with the chaperone GDI (guanine nucleotide displacement inhibitor) is needed. Structural and biophysical analysis of the Ypt1–GDI complex has revealed two components of this interaction relevant for Rab targeting (Pylypenko et al., 2006). Domain I of GDI interacts with the switch II region of Ypt1 only when this is in the GDP-bound inactive form. The doubly prenylated C terminus of Ypt1 occupies a hydrophobic cavity created by domain II of GDI. Simulation of this system and direct biophysical measurements suggests that in the absence of other factors GDI will rapidly deliver Rabs to and extract them from a lipid bilayer (Pylypenko et al., 2006; Wu et al., 2010). These ideas can be combined into a simple model for Rab activation at specific membrane surfaces (Fig. 1 A). In simple terms this model is a form of molecular speed-dating in which the Rab spends a short time sampling each membrane surface it encounters before finally meeting its cognate GEF partner, triggering a period of longer residence at that site (Fig. 1 A). In this model, GEF-mediated nucleotide exchange renders the Rab resistant to extraction by GDI, and thus drives accumulation of the active GTP-bound form of the Rab. This active Rab can then recruit effector proteins to the membrane surface and promote the desired recognition event. Such a system is analogous to the rapid proofreading of amino-acyl tRNAs during protein synthesis by the ribosome (Ibba and Söll, 1999). All amino-acyl tRNAs can enter the so-called acceptor site, but only if stable codon recognition occurs is the peptidyltransferase reaction initiated, otherwise the tRNA is rejected (Steitz, 2008). The two-stage kinetic proofreading of membrane surfaces by Rabs may similarly increase fidelity at little overall cost to the rate of vesicular traffic.Open in a separate windowFigure 1.The Rab activation and inactivation cycle. (A) Prenylated Rabs (black wavy lines) are bound by the chaperone GDI in the cytosol. Partitioning of the prenylated tail moiety between the hydrophobic pocket in GDI and the membrane bilayer allows Rabs to rapidly and reversibly sample membrane surfaces. When the GDP-bound inactive Rab encounters a cognate GEF nucleotide exchange occurs. This GTP-bound active Rab species does not interact with GDI and can therefore accumulate on the membrane surface, where it may further recruit effector proteins with specific biological functions. This cycle is reset when a GTP-bound Rab encounters a GAP (GTPase-activating protein) and the bound GTP is hydrolyzed to generate GDP and inorganic phosphate. (B) Additional specification of membrane domains within complex organelles, such as tubular domains of endosomes, or the fenestrated rims and different cisternae of the Golgi apparatus, may involve membrane receptors for Rabs (shown as light blue, dark blue, and green boxes). This could either involve (a) sequestration of the active Rab to a subdomain defined by the membrane receptor, or (b) direction of GDI unloading of an inactive Rab to specific sites on the organelle membrane also defined by a membrane receptor. Accumulation of a Rab at a specific site may be favored by GAPs opposing Rab activation at unwanted sites (Haas et al., 2007).Although this minimal Rab-targeting system does not require any additional factors, it is important to mention that this does not mean such factors do not exist. A family of membrane proteins with prenylated Rab-binding activity that can promote dissociation of some prenylated Rabs from GDI and favor retention of the GDP-bound form of the Rab downstream of membrane delivery by GDI has been identified (Dirac-Svejstrup et al., 1997; Martincic et al., 1997; Hutt et al., 2000; Sivars et al., 2003). These may therefore favor Rab activation, although recent data has suggested that such factors are not generally essential (Blümer et al., 2013). Intriguingly, other evidence links this family of proteins to factors involved in shaping subdomains of the ER and to the Golgi apparatus (Yang et al., 1998; Calero et al., 2001; Chen et al., 2004; Voeltz et al., 2006), perhaps suggesting that they may play roles in defining at which subdomain of an organelle an active Rab is enriched (Fig. 1 B).In addition to these regulatory factors, covalent modification can also be used to modulate the Rab activation cycle. Phosphorylation of Rab1 and Rab4 in mitosis alters the fraction of these GTPases that can associate with membranes (Bailly et al., 1991; van der Sluijs et al., 1992), although the exact mechanisms remain unclear. Furthermore, emerging evidence indicates that one Rab in yeast, Ypt11, is controlled by a phosphorylation-dependent mechanism regulating its activation and abundance (Lewandowska et al., 2013). A number of bacterial pathogens also encode enzymes that directly modify Rab GTPases and as a consequence alter the Rab regulatory cycle. During Legionella infection, Rab1 is modulated by a cycle of adenylylation and de-adenylylation by DrrA and SidA, respectively, and this modification of the conserved tyrosine residue in the switch II renders the protein constitutively active (Müller et al., 2010; Neunuebel et al., 2011; Tan and Luo, 2011). DrrA also has a GEF domain and can therefore directly activate and trap Rab1 in an active form independent of other cellular factors (Schoebel et al., 2009). A second bacterial protein, AnkX, mediates phosphocholination of an adjacent serine within the switch II region (Mukherjee et al., 2011; Campanacci et al., 2013). Pathogens such as Legionella use this covalent modification of Rabs to modulate their localization and activation (Stein et al., 2012). Although cellular enzymes that carry out related modification of Rabs are currently unknown, it would be premature to dismiss the possibility of their existence and use by cells to similarly control Rab activation and inactivation at specific sites.

Evidence for Rab activation on vesicle and target membrane surfaces

Based on the model and discussion so far it seems obvious that Rabs accumulate on the same membrane as their cognate GEF. Indeed, there is evidence that Rab1 may be activated and recruit the p115 tethering factor during the COP II vesicle formation stage of ER-to-Golgi transport (Allan et al., 2000). This would have the advantage that identity would be created at an early stage in vesicle biogenesis, and the vesicle could therefore be tethered to the Golgi before completion of the vesicle, thus increasing targeting efficiency. However, there is also evidence that Rab activation can occur at the target membrane and not only on a vesicle surface. Careful analysis of cell-free ER–Golgi transport assays revealed that Ypt1–Rab1 is not always required on the vesicle fraction, but is essential on the target Golgi membranes (Cao and Barlowe, 2000). Furthermore, a Ypt1 mutant with reduced nucleotide hydrolysis (which prevents its recycling from the Golgi compartment; Richardson et al., 1998), or Golgi membrane-anchored forms of Ypt1 (Cao and Barlowe, 2000) both support apparently normal ER–Golgi transport and cell growth. Subsequently, it was found that the COP II coat required to form ER–Golgi transport vesicles is the membrane receptor for the Ypt1–Rab1 GEF TRAPP (transport protein particle; Jones et al., 2000; Wang et al., 2000; Cai et al., 2007), indicating that Rab1 activation may occur on the coated vesicle. This raises questions about how the cytosolic Rab–GDI complex can access the membrane surface of a still-coated vesicle. However, because the COP II coat has an open lattice structure (Faini et al., 2013), it may be possible in this case for Ypt1–Rab1 to approach the membrane and insert. A further possibility is that COP II vesicles recruit TRAPP and promote the activation of Ypt1–Rab1 at the adjacent Golgi membranes to signal that an ER-derived vesicle is in close proximity (Fig. 2 A). This Golgi pool of activated Rab would then recruit effector proteins such as Uso1/p115 that trap and tether the incoming vesicle by directly engaging with vesicle SNAREs (Cao et al., 1998; Shorter et al., 2002).Open in a separate windowFigure 2.Recruitment mechanisms for Rab GEFs. Rab GEFs can be divided into two groups according to the mechanism of membrane recruitment. (A) Discrete coat protein complexes (green) recruit the first group. For example, COP II recruits the Rab1 GEF TRAPP to ER-Golgi vesicles, while clathrin-AP2 recruits DENND1A, the Rab35 GEF, to endocytic sites at the cell surface. In the case of TRAPP, biochemical and genetic data suggest that Rab1 can be activated on the target membrane, before vesicle tethering and SNARE-mediated fusion. (B) The larger second group of Rab GEFs is recruited by Rab GTPases either alone or in combination with a second factor (Rabs/factors listed next to arrow). For example, the GEF Sec2 is recruited to late-Golgi vesicles trafficking to the bud in yeast by the activated Rab Ypt31/32 and phosphatidylinositol 4-phosphate (PI4P), where it activates the Rab Sec4 (Rab8 in humans). The Rabex5–rabaptin complex, which is a Rab5 GEF, interacts with activated Rab4 or Rab5 and ubiquitylated cargo proteins on endocytic vesicles and early endosomes. A number of other GEFs (some additional examples shown) have been found to interact with active Rabs. Whether or not these represent the sole mode of membrane interaction for these GEFs is not defined at this time. PM, plasma membrane. (C) In situations where the GEF for a second Rab in the pathway is an effector for the first, a cascade can develop, where Rab-A promotes the recruitment of GEF-B for this second Rab-B.

Rab GEF targeting and regulation

The mechanism of GEF targeting is of crucial importance for understanding how Rabs are activated at a particular membrane site. At present, two different solutions to the problem depending on the GEF are known. First, as already mentioned, is vesicle coat–dependent GEF targeting (Fig. 2 A). Three examples are known at present: COP II recruitment of the Rab1 GEF TRAPP-I (Cai et al., 2007), and clathrin-adaptor protein complex 2–dependent recruitment of either the Rab35 GEF DENND1A (Allaire et al., 2010; Yoshimura et al., 2010) or the Rab5 GEF RME-6 (Sato et al., 2005; Semerdjieva et al., 2008) during endocytic transport from the plasma membrane. In the latter cases the exact nature of the membrane on which the target Rab is activated is unclear, but it is tempting to speculate that like COP II, the coated vesicle promotes Rab activation on the target organelle to signal the presence of an incoming vesicle to be tethered. The second larger group of GEFs comprises those known to interact with active Rab GTPases (Fig. 2 B). The first of these Rab GEF effectors defined was the Rabex-5–rabaptin complex, which is both a Rab5 exchange factor and effector for Rab4 and Rab5 (Horiuchi et al., 1997). Rabex-5 also binds to ubiquitin via a specific domain and this is important for regulating its recruitment to early endosomes (Lee et al., 2006; Mattera et al., 2006; Mattera and Bonifacino, 2008) where it activates Rab5.Specific phosphatidylinositols play a key role in defining membrane identity (Di Paolo and De Camilli, 2006)), and this is in part due to a role in recruitment or regulation of Rab exchange factors. Sec2, the exchange factor for Sec4–Rab8, is recruited to post-Golgi vesicles by a combination of the Rab Ypt32 and phosphatidylinositol 4-phosphate generated by Pik1 (Ortiz et al., 2002; Sciorra et al., 2005; Mizuno-Yamasaki et al., 2010). Similarly, in mammalian cells the Rab GEF Sec2–Rabin8 is recruited by the Ypt31/32 orthologue Rab11 (Knödler et al., 2010), and phosphatidylinositol 4-phosphate generated by the Pik1 orthologue Fwd is important for Rab11 regulation in Drosophila (Polevoy et al., 2009). Although less is known about the targeting of other Rab GEFs, the clear theme is that many are effectors for a Rab other than the one they activate (Fig. 2 B). The Ric1–Rgp1 complex is a GEF for Rab6 and effector for Rab33B at the Golgi (Pusapati et al., 2012) and the Rab21 GEF VARP is an effector for Rab32/38 (Zhang et al., 2006; Tamura et al., 2009). Additionally, a GEF for Rab32/38 is an effector for Rab9 (Kloer et al., 2010; Gerondopoulos et al., 2012), and the DENND5A Rab39 GEF is an effector for Rab6 (Recacha et al., 2009; Yoshimura et al., 2010). In addition to these canonical trafficking functions there are specialized examples that indicate there is some plasticity to both GEF targeting and specificity. The Ypt1 GEF TRAPP exists in an alternate form (TRAPP-II) with additional subunits that promote late-Golgi targeting and may create additional GEF activity toward Ypt31/32 (Morozova et al., 2006). Interestingly, in higher eukaryotes there is evidence that TRAPP-II may regulate the Ypt31/32 orthologues Rab11 in male meiotic cytokinesis in flies (Robinett et al., 2009) and Rab-A in plant cell polarization and division (Qi et al., 2011), respectively. TRS85 in another alternate TRAPP complex (TRAPP-III) promotes localization to the forming autophagosome and activates Rab1 during autophagy (Lynch-Day et al., 2010).The counterpart to this interlinked network of Rab activation is an equally complex set of interactions between Rabs and Rab GAPs. The GAP Gyp1 is an effector for Ypt32 and promotes GTP hydrolysis by Ypt1 in budding yeast (Rivera-Molina and Novick, 2009). In the absence of Gyp1, Ypt1 spreads into the later compartments of the secretory pathway that should be occupied by Ypt32 (Rivera-Molina and Novick, 2009). Interestingly, one of the cellular GAPs for Ypt1–Rab1 is a transmembrane protein of the ER that may prevent Rab1 activity from spreading earlier in the pathway to the ER rather than act to terminate Rab1 activity at the Golgi (Haas et al., 2007; Sklan et al., 2007). Similarly, two related proteins, RUTBC1 and RUTBC2, bind to active Rab9 and are GAPs for Rab32 and Rab36, respectively (Nottingham et al., 2011, 2012).Together, these findings have led to the general idea that the order of trafficking events in a pathway can potentially be defined by a series of Rabs acting as a cascade (Fig. 2 C). In such models one Rab triggers the next in the pathway by recruiting its cognate GEF, and then feedback develops as a GTPase-activating protein (GAP) is recruited to terminate the action of the previous Rab in the series (Mizuno-Yamasaki et al., 2012; Pfeffer, 2013). In part, this simply passes the problem on because we are then left with the question of how the previous Rab in the pathway or a cofactor for recruitment such as phosphatidylinositol 4-phosphate or ubiquitin is localized and generated only when required. In the case of the secretory pathway the ER provides a defined starting point where activation of Rab1–Ypt1 will inevitably result in a defined and correctly timed wave of Rab activation through the secretory pathway. However, a note of caution is needed when considering these ideas because far more support from experimental data looking at the biochemical properties of these systems both in vitro and in vivo is required to come to any definitive conclusions.

Ultrasensitive Rab activation switches

One of the key tenets of the membrane identity hypothesis is that Rabs should rapidly and accurately establish membrane identity and then be lost once the membrane recognition event is over. Although biochemical data on Rab GEFs clearly indicate these molecules generally have sufficiently high specificity to ensure activation of only one Rab or a set of closely related Rabs (Delprato et al., 2004; Yoshimura et al., 2010; Gerondopoulos et al., 2012), how rapid switch-like accumulation is ensured is less obvious. Similar issues exist for termination of the Rab cycle by Rab GAPs. As already mentioned, Rab cascade models give part of the solution to this problem, and provide features that can ensure vectorial flow in a membrane traffic pathway (Mizuno-Yamasaki et al., 2012; Pfeffer, 2013). However, they do not fully explain how switch-like transitions and defined compartmental boundaries are achieved (Del Conte-Zerial et al., 2008). A possible solution to this problem comes from studies on the regulation of other complex biological systems, exemplified by control of cell cycle transitions (Tyson et al., 2001). Rather than displaying the expected Michaelis-Menten kinetics (Fig. 3 A), Rab cycles may yield properties of ultrasensitivity (Goldbeter and Koshland, 1981, 1984). This would appear to be a valid proposal if the Rab cycle is treated as being analogous to a covalent modification (Rab and Rab-modified, for GDP and GTP forms, respectively) and because GEF activity is generally assumed to be limiting (Blümer et al., 2013). In such a situation, inputs activating the GEF, for example membrane recruitment requiring multiple or binding of an activator, would be amplified and give rise to very large changes in the amount of activated Rab (Fig. 3 A). When combined with feedback loops, this can create a bistable switch between two states as shown for cell cycle transitions (Novak and Tyson, 1993; Pomerening et al., 2003). In the case of GTPase regulation, as the input controlling the GEF increases then the system transitions to a Rab-active state that remains stable over a wide range of GEF activity. GAP activation could then trigger exit from this state. This is also useful for providing a potential explanation for the timing properties of a Rab cascade. Ultrasensitivity and bistability are therefore likely to be useful concepts when explaining the behavior of Rabs, especially when considering complex interlinked cycles (Fig. 3 B) because they avoid the futile cycles where GAPs and GEFs fight one another and thus don’t do any useful work.Open in a separate windowFigure 3.Ultrasensitivity and bistability in Rab regulatory networks. (A) A simplified schematic of a Rab activation cycle is shown treating GDP–GTP exchange as equivalent to a covalent modification cycle such as phosphorylation. Because the reaction can only occur at a membrane surface, membrane recruitment factors are treated as activating inputs. Assuming no feedback and normal first-order reaction kinetics, Rab recruitment would be expected to follow Michaelis-Menten behavior. In cases where substrate is saturating and the reaction becomes zero-order, Goldbeter and Koshland (1984) have shown that product formation becomes more sensitive to enzyme concentration. In this case, generation of GTP-bound Rab becomes ultrasensitive to GEF concentration at the membrane surface. If additional positive feedback controls exist as shown in the bottom panel, then bistability may develop. In this case a rapid switch-like transition in Rab activity develops as Rab GEF concentration increases. Once in the active state the system becomes less dependent on continued high GEF activity. (B) A model for an interlinked Rab cascade is shown. The GEF for Rab-B is an effector for activated Rab-A, while the GAP for Rab-A is regulated by Rab-B. An example of this latter situation is provided by the Ypt1–Yp32 system discussed in the main text and shown in the bottom panel, where a Ypt1 GAP Gyp1 is an effector for Ypt32 (Rivera-Molina and Novick, 2009) and inhibits Ypt1. This coupling of the two cycles can result in coupled ultrasensitive switch-like transitions or bistability.A groundbreaking study in this area has applied these ideas to the conversion of Rab5-positive early endosomes to Rab7-positive late endosomes and lysosomes (Del Conte-Zerial et al., 2008). This analysis has provided strong evidence that positive and negative feedback loops in this system mediated by Rab GEFs and GAPs result in bistability in the form of a cut-out switch, so that Rab5 accumulation is followed by an abrupt transition at which Rab5 is rapidly lost and Rab7 accumulates (Del Conte-Zerial et al., 2008). Underpinning this is a biochemical network in which the Mon1–Ccz1 Rab7 GEF complex displaces Rabex-5, thus breaking the positive feedback loop to Rab5 activation (Poteryaev et al., 2010) and simultaneously promoting recruitment and activation of Rab7 (Nordmann et al., 2010; Gerondopoulos et al., 2012). Although there are only few studies where these ideas have been considered, they can be experimentally tested and are likely to be of increasing importance in membrane traffic regulation.

Origins of Rab GTPase control systems

One of the most difficult questions in membrane trafficking relates to the origins of complex internal membrane systems in eukaryotes. Analysis of Rab GTPases themselves suggests a pattern of evolution of Rabs consistent with the evolution of a core set of membrane organelles of the endocytic and secretory pathways (Diekmann et al., 2011; Klöpper et al., 2012). Yet, this provides little insight into how membrane organelles initially arose. Recent data on the structure of Rab GTPase regulators and coat protein complexes has identified common features with GTPase regulators in other systems including prokaryotes (Kinch and Grishin, 2006; Zhang et al., 2012; Levine et al., 2013). The conserved Longin–Roadblock fold has emerged as a structural feature of the large family of DENN-domain Rab GEFs in human cells (Yoshimura et al., 2010; Wu et al., 2011; Levine et al., 2013). Intriguingly, related domains are also present in the signal sequence receptor involved in protein translocation into the ER, vesicle coat protein complexes, and the MglA GTPase–MglB bacterial cell polarity regulator (Sun et al., 2007; Miertzschke et al., 2011; Levine et al., 2013). Although far from conclusive, these findings provide important pointers to the development of GTPase control systems, and more generally the early origins of membrane traffic pathways in eukaryotes from membrane-associated GTPases and their effector proteins.Are Rabs alone capable of triggering the pathways defining membrane identity? Multiple lines of evidence show Rab GTPases are clearly important and far from inconsequential regulators of vesicle traffic; however, further evidence is required before we should conclude that they are causal regulators of vesicle or organelle membrane identity. Neither of the studies using strategies to modulate the cellular localization of Rab GEFs reported that the mitochondria altered their identity or were converted into an endosome or Golgi because of the mistargeted Rabs (Gerondopoulos et al., 2012; Blümer et al., 2013). The picture emerging is therefore one in which Rabs cannot program membrane identity alone and must work in concert with other factors. Defining and reconstituting the systems needed to create membrane identity is therefore a major goal for membrane traffic research.     

The Presence of Fucogalactoxyloglucan and Its Synthesis in Rice Indicates Conserved Functional Importance in Plants

The predominant structure of the hemicellulose xyloglucan (XyG) found in the cell walls of dicots is a fucogalactoXyG with an XXXG core motif, whereas in the Poaceae (grasses and cereals), the structure of XyG is less xylosylated (XXGG_n core motif) and lacks fucosyl residues. However, specialized tissues of rice (Oryza sativa) also contain fucogalactoXyG. Orthologous genes of the fucogalactoXyG biosynthetic machinery of Arabidopsis (Arabidopsis thaliana) are present in the rice genome. Expression of these rice genes, including fucosyl-, galactosyl-, and acetyltransferases, in the corresponding Arabidopsis mutants confirmed their activity and substrate specificity, indicating that plants in the Poaceae family have the ability to synthesize fucogalactoXyG in vivo. The data presented here provide support for a functional conservation of XyG structure in higher plants.The plant cell wall protects and structurally supports plant cells. The wall consists of a variety of polymers, including polysaccharides, the polyphenol lignin, and glycoproteins. One of the major polysaccharides present in the primary walls (i.e. walls of growing cells) in dicots is xyloglucan (XyG), which consists of a β-1,4-glucan backbone with xylosyl substituents. XyG binds noncovalently to cellulose microfibrils and thereby, is thought to act as a spacer molecule, hindering cellulose microfibrils to aggregate (Carpita and Gibeaut, 1993; Pauly et al., 1999a; Bootten et al., 2004; Cosgrove, 2005; Hayashi and Kaida, 2011; Park and Cosgrove, 2012).The side-chain substitutions on XyG can be structurally diverse depending on plant species, tissue type, and developmental stage of the tissue (Pauly et al., 2001; Hoffman et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009, 2012; Lampugnani et al., 2013; Schultink et al., 2014). A one-letter code nomenclature has been established to specify the XyG side-chain substitutions (Fry et al., 1993; Tuomivaara et al., 20145). According to this nomenclature, an unsubstituted glucosyl residue is indicated by a G, whereas a glucosyl residue substituted with a xylosyl moiety is shown as an X. In most dicots, the xylosyl residue can be further substituted with a galactosyl residue (L), which in turn, can be further decorated with a fucosyl residue (F) and/or an acetyl group (F/L). In some species, the xylosyl residue can be substituted with an arabinosyl moiety (S), and the backbone glucosyl residue can be O-acetylated (G; Jia et al., 2003; Hoffman et al., 2005).Numerous genes have been identified in Arabidopsis (Arabidopsis thaliana) that are involved in fucogalactoXyG biosynthesis (Fig. 1; Pauly et al., 2013; Schultink et al., 2014). The glucan backbone is thought to be synthesized by cellulose synthase-like C (CSLC) family proteins, such as AtCSLC4, as shown by in vitro activity data (Cocuron et al., 2007). Several xylosyltransferases (XXTs) from glycosyl transferase family 34 (GT34) are thought to be responsible for XyG xylosylation. Five of these XXTs in Arabidopsis seem to have XXT activity on XyG in vitro (Faik et al., 2002; Zabotina et al., 2008; Vuttipongchaikij et al., 2012; Mansoori et al., 2015). MURUS3 (MUR3) represents a galactosyltransferase that transfers galactosyl moieties specifically to xylosyl residues adjacent to an unsubstituted glucosyl residue on an XXXG unit, converting it to XXLG, whereas Xyloglucan L-side chain galactosyl Transferase2 (XLT2) was identified as another galactosyltransferase transferring a galactosyl moiety specifically to the second xylosyl residue, resulting in XLXG (Madson et al., 2003; Jensen et al., 2012). Both MUR3 and XLT2 belong to GT47 (Li et al., 2004). MUR2/FUCOSYLTRANSFERASE1 (FUT1) from GT37 was found to harbor fucosyltransferase activity, transferring Fuc from GDP-Fuc to a galactosyl residue adjacent to the unsubstituted glucosyl residue (i.e. onto XXLG but not onto XLXG; Perrin et al., 1999; Vanzin et al., 2002). O-acetylation of the galactosyl residue is mediated by Altered Xyloglucan4 (AXY4) and AXY4L, both of which belong to the Trichome Birefringence-Like (TBL) protein family (Bischoff et al., 2010; Gille et al., 2011; Gille and Pauly, 2012).Open in a separate windowFigure 1.Schematic structures of two types of XyGs and known biosynthetic proteins in Arabidopsis (Hsieh and Harris, 2009; Pauly et al., 2013). The corresponding one-letter code for XyG is shown below the pictograms (Fry et al., 1993; Tuomivaara et al., 2015).XyG found throughout land plants exhibits structural diversity with respect to side-chain substitution patterns (Schultink et al., 2014). Most dicots, such as Arabidopsis, and the noncommelinoid monocots possess a fucogalactoXyG of the XXXG-type XyG structure as shown in Figure 1. However, plant species in the Solanaceae and Poaceae as well as the moss Physcomitrella patens contain a different XyG structure with a reduced level of xylosylation, resulting in an XXGG_n core motif (York et al., 1996; Kato et al., 2004; Gibeaut et al., 2005; Jia et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009). In addition, the glucan backbone can be O-acetylated in plants of Solanaceae and Poaceae families (Gibeaut et al., 2005; Jia et al., 2005). XyG from Solanaceae with an XXGG core motif can be further arabinosylated and/or galactosylated (Jia et al., 2005). No XyGs with an XXGG_n motif backbone have been reported to be fucosylated.The function of structural diversity of XyG substitutions, such as fucosylation and/or altered xylosylation pattern, remains enigmatic. Removing the terminal fucosyl or acetyl moieties in the corresponding Arabidopsis mutants does not lead to any change in plant growth and development (Vanzin et al., 2002; Gille et al., 2011). However, removing galactosyl residues as well as fucosyl and acetyl moieties in the Arabidopsis xlt2 mur3.1 double mutant results in a dwarfed plant (Jensen et al., 2012; Kong et al., 2015). Replacing the galactosyl moiety with an arabinofuranosyl residue by, for example, expressing a tomato (Solanum lycopersicum) arabinosyltransferase in the Arabidopsis xlt2 mur3.1 mutant rescues the growth phenotype and restores wall biomechanics, indicating that galactosylation and arabinosylation in XyG have an equivalent function (Schultink et al., 2013). Recently, fucosylated XyG structures were found in the pollen tubes of tobacco (Nicotiana alata) and tomato, indicating that fucogalactoXyG is likely also present in other Solanaceae plants, albeit restricted to specific tissues (Lampugnani et al., 2013; Dardelle et al., 2015). Although there is circumstantial evidence that fucogalactoXyG is present in cell suspension cultures of rice (Oryza sativa) and cell suspension cultures of fescue (Festuca arundinaceae; McDougall and Fry, 1994; Peña et al., 2008), fucogalactoXyG has not been found in any physiologically relevant plant tissues of members of the Poaceae (Kato et al., 1982; Watanabe et al., 1984; Gibeaut et al., 2005; Hsieh and Harris, 2009; Brennan and Harris, 2011). Here, we provide chemical and genetic evidence that fucogalactoXyG is, indeed, present in plant tissues of a grass (rice) and prove that the rice genome harbors the genes that could be part of the synthetic machinery necessary to produce fucogalactoXyG.