Cell biology in development: The cell biology of planar cell polarity

Planar cell polarity (PCP) refers to the coordinated alignment of cell polarity across the tissue plane. Key to the establishment of PCP is asymmetric partitioning of cortical PCP components and intercellular communication to coordinate polarity between neighboring cells. Recent progress has been made toward understanding how protein transport, endocytosis, and intercellular interactions contribute to asymmetric PCP protein localization. Additionally, the functions of gradients and mechanical forces as global cues that bias PCP orientation are beginning to be elucidated. Together, these findings are shedding light on how global cues integrate with local cell interactions to organize cellular polarity at the tissue level.The collective alignment of cell polarity across the tissue plane is a phenomenon known as planar cell polarity (PCP). Exemplified by the uniform orientation of bristles covering the insect epidermis or of the hairs covering the mammalian body surface (Fig. 1 A), PCP patterns can align over thousands, even billions of cells. This phenomenon is controlled by the so-called PCP pathway, which integrates global directional cues to produce locally polarized cell behaviors. There has been a recent surge in interest in PCP after discoveries that various processes such as vertebrate gastrulation, mammalian ear patterning and hearing, and neural tube closure all require a conserved set of PCP genes (Heisenberg et al., 2000; Tada and Smith, 2000; Wallingford et al., 2000; Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Montcouquiol et al., 2003; Copley et al., 2013). Since that time, the PCP pathway has been found to coordinate cell behaviors in numerous diverse settings including polarized ciliary beating in the trachea and brain ventricles (Tissir et al., 2010; Vladar et al., 2012), oriented cell divisions (Gong et al., 2004; Baena-López et al., 2005; Ségalen et al., 2010; Mao et al., 2011), lung branching (Yates et al., 2010), and hair follicle alignment (Guo et al., 2004; Devenport and Fuchs, 2008), to name a few (Fig. 1). Genetic disruptions to PCP cause severe developmental abnormalities in vertebrates, notably neural tube defects, left/right patterning defects, and ciliopathies, which highlights the essential requirement for PCP in development (Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Wang et al., 2006a,b; Kim et al., 2010; Song et al., 2010).Open in a separate windowFigure 1.Planar cell polarity and the core PCP components. (A and B) The Drosophila wing blade and mammalian epidermis illustrate the phenomenon of PCP. In both cases, hairs point in a single direction along the tissue axis, where they align locally with neighboring hairs and globally across the tissue. Whereas Drosophila wing hairs are produced by single cells, mammalian hairs emerge from multicellular hair follicles, which orient as a unit. A conserved PCP pathway controls the collective alignment of both types of structures. (C) Core PCP components localize to the plasma membrane and asymmetrically segregate along the epithelial plane as indicated.Like many types of cell polarity, the establishment of PCP involves (1) a global orienting cue, (2) asymmetric segregation of dedicated polarity proteins, and (3) translation of polarity information into polarized outputs. But unlike other types of cell polarity, the PCP mechanisms we currently understand involve coupling between adjacent cells, allowing for the alignment of polarity over many cell distances.First described in insects and then genetically dissected in Drosophila melanogaster, PCP was long confined to the realm of experimental embryology and genetics until the discovery that the protein products of several PCP genes were localized asymmetrically within the cell, thrusting PCP into the domain of cell biology (for review see Strutt and Strutt, 2009). The challenge to understanding PCP on a molecular level is that long-range PCP is, in essence, an in vivo phenomenon that is difficult to recapitulate in a tissue culture dish. However, recent advances in imaging technology combined with increasingly sophisticated genetic tools are helping us to decipher the in vivo cell biology of PCP. In this review, I highlight some of the recent advances made toward understanding the cell biology underlying the establishment of coordinated polarized cell behaviors. For clarity, I limit discussions of PCP phenomena that meet the definition of PCP proposed by Goodrich and Strutt (2011): namely, that “cell–cell communication causes two or more cells to adopt coordinated polarity” in a process that is mechanistically “dependent upon planar polarity proteins.” Other aligned cellular patterns or examples of noncanonical Wnt signaling, sometimes described as “Wnt/PCP” signaling, will not be discussed.

PCP components and molecular asymmetries

Two molecular systems control PCP behavior, the “core” and the “Fat–Dachsous (Ft–Ds)” PCP pathways. A key feature of both is the asymmetric distribution of their constituents (Fig. 2). The core PCP pathway is composed of the multipass transmembrane proteins Frizzled (Fz), Van Gogh (Vang; also known as Strabismus/Stbm), and Flamingo (Fmi; also known as Starry night/Stan), and the cytosolic components Dishevelled (Dsh), Prickle (Pk), and Diego (Dgo). On one edge of the cell reside Fz, Dsh, and Dgo, and on the opposite side lie Vang and Pk (Figs. 1 C and 2 B; Axelrod, 2001; Strutt, 2001; Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). The atypical cadherin, Fmi, resides on both sides, where it forms homodimers between neighboring cells (Usui et al., 1999; Shimada et al., 2001). These molecular asymmetries are observed in sensory hair cells of the vertebrate inner ear (Wang et al., 2005, 2006a,b; Montcouquiol et al., 2006; Deans et al., 2007; Song et al., 2010), the mammalian epidermis (Devenport and Fuchs, 2008; Devenport et al., 2011), brain ventricles (Tissir et al., 2010), and trachea (Vladar et al., 2012). Mutations in core PCP components lead to a loss or randomization of polarity and misalignment of cellular structures along the tissues axis.Open in a separate windowFigure 2.Asymmetric localization of PCP components. (A) PCP asymmetry develops progressively from an initially uniform distribution of core PCP proteins. Fz, Dsh, and Dgo (red) localize to the distal/posterior edge, whereas Vang and Pk (turquoise) localize to the proximal/anterior side. Fmi (dark blue) localizes to both sides, where it forms homodimers between neighboring cells. (B) Feedback interactions between core PCP components. A Fz–Fmi complex interacts preferentially with a Vang–Fmi complex between cells, whereas proximal and distal complexes antagonize one another within the cell. (C) Ds and Fj are expressed in opposing gradients in the Drosophila wing blade. Fj positively modulates Ft activity, leading to a gradient of Ft activity across the wing (not depicted). (D) Graded expression of Ds and Fj leads to asymmetric cellular localization of Ds and Ft, which form heterodimers between adjacent cells. Dachs, a downstream component of the Ft–Ds pathway, also localizes asymmetrically in association with Ds.The Ft–Ds pathway includes the large protocadherins Ft and Ds and the Golgi resident transmembrane kinase, Four-jointed (Fj; for review see Matis and Axelrod, 2013; Thomas and Strutt, 2012). Similar to the core system, Ft–Ds also displays molecular asymmetries in flies. Ds and its ligand Ft accumulate on opposite cell edges, where they form intercellular heterophilic interactions (Fig. 2 D; Matakatsu and Blair 2004; Ambegaonkar et al., 2012; Brittle et al., 2012). Unlike the core components, Ds and Fj are expressed in complementary gradients in the Drosophila eye and developing wing, which contribute to the cellular asymmetries of Ds and Ft (Fig. 2, C and D). Whether Ft–Ds–Fj gradients and asymmetries are conserved in vertebrate systems has yet to be determined.

Segregation of cortical polarity proteins: Shaking hands with the enemy

The asymmetric segregation of Fz–Dsh–Fmi and Vang–Pk–Fmi complexes to opposite sides of the cell relies on their mutual exclusion intracellularly and their preferential binding between neighboring cells (Fig. 2 B; for review see Strutt and Strutt, 2009). There is mutual interdependence among core PCP components for their asymmetric localization. Depletion of any one core PCP component results in a loss of asymmetry of all the others. In addition, PCP asymmetry develops progressively from initially uniform distributions (Fig. 2 A). Thus, PCP asymmetry can be thought of not as a simple hierarchy of interactions, but the result of feedback amplification of an initial directional bias.

Intercellular interactions.

PCP requires cell–cell communication, mediated by the transmembrane components of the core system, where it is thought that Fz–Fmi on one cell interacts with Vang–Fmi on its neighbor. These interactions are best understood in the Drosophila wing blade, where PCP controls the alignment of wing hairs along the proximal–distal axis (Figs. 1 A and 2, A and B). In the wing, Vang–Pk localize to the proximal face of each cell, whereas Fz–Dsh–Dgo localize distally adjacent to the wing hair (Figs. 1 C and 2, A and B; Axelrod, 2001; Strutt, 2001; Tree et al., 2002; Bastock et al., 2003; Das et al., 2004). By generating mutant clones and examining PCP localization at the clone border, the intercellular interactions between neighboring cells can be assessed in vivo. For example, when Fz is lacking within a clone, leaving only Vang–Fmi available at cell junctions, then Fz–Fmi in adjacent wild-type cells is recruited to the clone border (Chen et al., 2008). Vang mutant clones produce a similar effect, but in this case the excess Fz recruits Vang to clone borders (Bastock et al., 2003). What mediates these intercellular asymmetric interactions? One possibility is that Vang and Fz interact directly, and in vitro binding assays between the Fz extracellular domain and Vang suggest that this mechanism is possible (Wu and Mlodzik, 2008). However, mutants of Fz or Vang lacking their extracellular domains can still recruit one another between cells, which suggests that something else must bridge the two proteins (Chen et al., 2008). The seven-pass transmembrane cadherin, Fmi, likely performs this function. Fmi is essential for the junctional recruitment of Fz and Vang, and Fmi homodimers appear to be functionally asymmetric (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012). Clonal overexpression of Fmi preferentially recruits Fz to the clone border, even in the absence of Vang, which suggests that excess or unpaired Fmi is in a configuration that has higher affinity for Fmi–Fz than Fmi–Vang (Chen et al., 2008; Strutt and Strutt, 2008). Thus, Fmi may exist in two forms depending on whether it is paired with Fz or Vang, but the molecular basis for this difference is not known (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012).

Amplification of asymmetry.

Intercellular Fz–Fmi and Vang–Fmi complexes can form between cells in any orientation, so how do they resolve into discrete and opposed asymmetric domains? One way is through clustering of Fz–Fmi and Vang–Fmi complexes of the same orientation, and the cytoplasmic PCP components are particularly important for this function. As PCP complexes grow increasingly asymmetric, they cluster into discrete puncta that are stably associated with the plasma membrane and are resistant to endocytosis (Strutt et al., 2011). FRAP analysis of Fz-containing puncta demonstrated that they are highly stable compared with diffuse Fz-GFP, and have limited lateral mobility within the membrane. In the absence of Dsh, Pk, or Dgo, the size, intensity, and stability of Fz-containing puncta are diminished (Strutt et al., 2011), whereas overexpression causes Fz accumulation and coalescence into larger puncta (Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). Although the precise mechanisms driving PCP puncta formation are not known, the cytoplasmic components do not affect endocytosis, which suggests that they contribute to puncta formation by clustering intercellular complexes (Strutt et al., 2011). Pk can interact homophilically (Jenny et al., 2003; Ayukawa et al., 2014), which might promote clustering of proximal Vang–Pk–Fmi complexes. It will also be interesting to determine whether the cytoskeleton is directly connected to PCP complexes to minimize lateral mobility within the membrane.A second mechanism contributing to PCP asymmetry is directed transport. Live imaging of fluorescently tagged PCP proteins in pupal wings showed that Fz- and Dsh-containing particles travel across the cell in a proximal-to-distal direction (Shimada et al., 2006; Matis et al., 2014; Olofsson et al., 2014). These particles most likely represent endosomes undergoing transcytosis, as they arise from the proximal cortex and are labeled by the endocytic tracer FM4-64. This mechanism could serve to amplify asymmetry or even provide the initial polarity bias by removing proximal Fz–Dsh–Fmi complexes and transporting them to the distal side. Directed PCP transport is mediated by an array of subapical, noncentrosomal microtubules (MTs) that align along the proximal–distal axis, with the plus ends oriented with a slight distal bias (Hannus et al., 2002; Shimada et al., 2006; Harumoto et al., 2010; Matis et al., 2014; Olofsson et al., 2014). Ft and Ds are required for proximal–distal MT alignment (Harumoto et al., 2010), which suggests that the Ft-Ds system may feed into the core PCP system by orienting cytoskeletal architecture to deliver Fz–Dsh–Fmi complexes to the distal edge of the cell.Directed transport of Vang-containing endosomes has not been reported in flies, but selective trafficking could target Vang to specific membrane domains. In mammalian cells, exit of the Vang homologue Vangl2 from the trans-Golgi network (TGN) requires Arfrp1 (an Arf-like GTPase) and the clathrin adaptor complex AP-1, neither of which are required for the transport of a mammalian Fz homologue Fz6 or Fmi/Celsr1, which suggests that the differential sorting of PCP complexes to opposite sides of the cell could initiate at TGN export (Guo et al., 2013). Whether newly synthesized Vang and Fz proteins are transported to opposing cell surfaces from the TGN has not yet been explored.Microtubule orientation also correlates with PCP asymmetry in mouse trachea epithelial cells, where PCP coordinates the alignment of motile cilia (Vladar et al., 2012). MTs are planar polarized with their plus ends oriented toward the Fz–Dvl domain, and disruption of MTs with nocodazole impairs core PCP localization. Similarly, MTs are needed to establish Pk asymmetry in gastrulating zebrafish embryos (Sepich et al., 2011). However, in the skin epithelium, MTs align perpendicular to the axis of PCP asymmetry (unpublished data). Thus, directed transport along MTs may not be required in all tissue types for the establishment of PCP asymmetry.

Negative regulation.

Repulsive interactions between Vang- and Fz-containing complexes may also contribute to the amplification of asymmetry, and cytoplasmic proteins have been proposed to perform this function. Pk and Dgo both bind to Dsh in vitro, interacting with the same domain on Dsh in a mutually exclusive manner (Jenny et al., 2005). In addition, overexpression of Pk can prevent Dsh translocation to the membrane (Tree et al., 2002; Carreira-Barbosa et al., 2003), which suggests that Pk binding to Dsh could displace it from the proximal side of the cell. On the distal side, Dgo binding to Dsh would prevent association with Pk, thus enhancing Dsh distal localization. This increase in Dsh and Pk asymmetry would then positively feed back by clustering the transmembrane components into stable membrane domains.Modulation of PCP protein levels by ubiquitin-mediated degradation also leads to feedback by restricting the amount of one PCP protein to antagonize another. In flies, regulation of Dsh by a Cullin-3-BTB E3 ubiquitin ligase complex limits its levels at cell junctions (Strutt et al., 2013a). Reduction of Cullin-3 leads to an increase in overall core PCP protein levels, a reduction of asymmetry, and defects in wing hair polarity, which is consistent with Dsh overexpression phenotypes (Strutt et al., 2013a). SkpA, a subunit of the SCF E3 ligase, regulates Pk levels by promoting its degradation in a Vang-dependent manner (Strutt et al., 2013b). In mice, Smurf E3 ligases ubiquitinate Pk and promote its local degradation by binding to phosphorylated Dvl2 (a mammalian homologue of Dsh; Narimatsu et al., 2009). Smurfs are required for Pk localization in the inner ear and floor plate, and their removal leads to defects in convergent extension (CE) and stereocilia alignment (Narimatsu et al., 2009). Thus, targeting Pk for degradation either balances total Pk protein levels or targets a specific pool of Pk for ubiquitination and proteasome degradation.

Tissue-level polarity cues: This way or that?

What provides the tissue-level polarity cue that biases core PCP asymmetry in one direction over another? This is perhaps the most fundamental, yet poorly understood, element of PCP. Current models propose that an upstream, graded cue provides an initial bias in PCP asymmetry by regulating the levels, localization, or activity of one or more of the core proteins. Gradients are attractive candidates for providing global polarity cues, as they can act across many cells and define the tissue boundaries over which polarity must be oriented.

Ft–Ds–Fj.

Unlike the core proteins, Ds and Fj are nonuniformly expressed in the Drosophila eye, wing, and abdominal segments, and as such, the Ft–Ds module has been proposed to provide a global polarity cue (Fig. 2, C and D; for review see Ma et al., 2003; Yang et al., 2002; Thomas and Strutt, 2012; Matis and Axelrod, 2013). Ft and Ds are heterodimeric cadherins, regulated by the Golgi kinase Fj (Ishikawa et al., 2008; Brittle et al., 2010; Simon et al., 2010). The complementary expression patterns of Ds and Fj are thought to give rise to asymmetric Ft and Ds protein localization, with Ft and Ds localizing to opposite sides of each cell (Fig. 2 D; Ambegaonkar et al., 2012; Bosveld et al., 2012; Brittle et al., 2012). Because Fj positively regulates the activity of Ft, a gradient of Ft activity is expressed across the wing complementary to that of its ligand, Ds (Simon et al., 2010). Mutations in the Ft–Ds system give rise to swirling wing hair patterns, and disrupt the global alignment of core PCP proteins, but not their asymmetric distributions.An appealing model for symmetry breaking in the early Drosophila wing is that cellular asymmetries of Ft–Ds polarize MT organization and promote the distal transport of Fz–Dsh–Fmi vesicles (Shimada et al., 2001, Harumoto et al., 2010; Matis and Axelrod, 2013; Matis et al., 2014). This would produce an initial bias in Fz–Dsh localization, which would then be amplified by feedback interactions. However, several pieces of evidence have prevented the model from gaining universal acceptance. First, Ds and Fj gradients are oriented in opposite directions with respect to the core PCP proteins in the wing compared with the eye and abdomen. This discrepancy has been rectified with the finding by two independent groups that cells interpret Ft–Ds–Fj gradients differently depending on which of two Pk isoforms is expressed (Ayukawa et al., 2014; Olofsson et al., 2014). Second, the Ft–Ds system can orient PCP independently of the core pathway, and thus the two systems orient polarity in parallel, as opposed to in a single, common pathway (Casal et al., 2006). Third, Ft–Ds mutations affect core PCP orientation only regionally in the wing, which suggests that, if Ft–Ds provides a global bias, other, redundant cues must also exist (Matakatsu and Blair, 2006; Matis et. al., 2014). Finally, the direction of Ft–Ds and core PCP asymmetry diverges late in wing development, where the two systems become completely uncoupled. Intriguingly, the extent of coupling depends on which isoform of Pk is expressed (Merkel et al., 2014). Perhaps the simplest explanation for Ft–Ds function is that it can both transmit polarity information independent of the core system and organize the cytoskeleton to provide an initial bias of core PCP asymmetry, but which mechanism predominates depends on the tissue and developmental stage.

Wnts.

Wnt proteins have long been considered attractive candidates to provide tissue-level polarity cues because Fz and Dsh are primary components of the Wnt–β-catenin signaling pathway. Wnts are secreted glycoproteins that bind to Fz and other receptors, and often display graded expression. In vertebrates, Wnts are clearly important regulators of PCP, but whether they act instructively or permissively is unclear. In zebrafish, Wnt5a and Wnt11 are required for CE movements during gastrulation, but uniform expression of Wnt11 rescues the mutant phenotype, which suggests that it is permissive rather than instructive (Heisenberg et al., 2000; Kilian et al., 2003). Wnt5a is expressed in a gradient along the axis of polarity in the mouse inner ear, where it interacts genetically with Vangl2 in cochlear hair cell orientation (Qian et al., 2007). In the mouse limb, Wnt5a and its atypical receptor Ror2 are required for limb elongation and the asymmetric localization of Vangl2 at the proximal face of converging and extending chondrocytes (Gao et al., 2011). Wnt5a is expressed in a distal-to-proximal gradient, which induces a gradient of Vangl2 phosphorylation. The functional consequences of Vangl2 phosphorylation are unknown but Vangl2 cellular asymmetry appears to be strongest distally, where Wnt5a and Vangl2 phosphorylation levels are highest (Gao et al., 2011).While several studies had argued against the involvement of Wnt proteins in Drosophila PCP (Lawrence et al., 2002; Chen et al., 2008), it was recently discovered that Wingless (Wg) and Wnt4a act redundantly to orient PCP in the wing, particularly near the wing margin (Wu et al., 2013). Misexpression of Wg or Wnt4a reorients wing hair polarity in a pattern reminiscent of Fz loss of function, which suggests that Wnt gradients may orient polarity by antagonizing Fz. Consistently, the ability of Fz and Vang to recruit one another between adjacent cells in culture was inhibited by the addition of Wg or Wnt4a, which suggests that Wnts could provide a polarizing cue by diminishing Fz–Vang interactions at the margin of the wing, where Wnt expression is highest (Wu et al., 2013). However, Wnt4a overexpression also reorients MT alignment, suggesting that Wnts may act as polarity cues thorough an effect on the cytoskeleton (Matis et. al., 2014). Alternatively, Sagner et al. (2012) suggest that Wg orients core PCP indirectly through its effects on wing patterning and growth. Although the evidence for Wnt gradients as global PCP cues is accumulating, the mechanisms by which they regulate core protein levels or activity remain to be elucidated.

Mechanical forces.

Anisotropic mechanical forces that accompany growth and morphogenesis can also provide global polarizing cues. During wing development, PCP reorients in response to extensive morphogenetic changes that elongate the wing along the proximal–distal axis. In early pupal wings, PCP aligns toward the wing margin and then reorients during wing elongation and contraction of the wing hinge (Aigouy et al., 2010). These morphogenetic changes have broad effects on cell behavior, inducing cell elongation, oriented divisions, and cell rearrangements with a concomitant reorientation of PCP. Severing the wing pouch from the hinge blocks cell flows and PCP reorientation, which suggests that the anisotropic tension from hinge contraction drives tissue flow and the reorientation of polarity (Aigouy et al., 2010). Although this model doesn’t explain what initially biases PCP, it does demonstrate how the morphogenetic processes that shape tissues can completely remodel global PCP alignment. This is an attractive model to explain how PCP aligns over very large tissues, like the mammalian skin, where hairs consistently reorient along regions of extensive tissue elongation such as the face, limbs, and ears.

Downstream effectors of PCP: Steering the wheel

If PCP is the cell’s compass, it is also the steering wheel, directing downstream, polarized cell behaviors in response to global directional cues. PCP can polarize a wide range of cell behaviors, which suggests that it can intersect with numerous downstream effectors. We focus here on three examples where the molecular mechanisms linking core PCP to their polarized outputs have recently been elucidated.

Distal positioning of wing hairs.

Each cell of the Drosophila wing blade emits a single actin-rich protrusion from its distal edge. The placement of the wing hair strongly correlates with the position of Fz–Dsh–Fmi, which suggests that core proteins may localize cytoskeletal regulators to distinct positions within the cell (Strutt and Warrington, 2008). On the proximal side, Vang recruits a group of proteins that negatively regulate actin prehair formation: Inturned, Fuzzy, and Fritz (Adler et al., 2004; Strutt and Warrington, 2008). These three proteins regulate Multiple Wing Hairs, a GTPase-binding/formin-homology 3 (GBD/FH3) domain protein thought to repress actin polymerization (Strutt and Warrington, 2008; Yan et al., 2008). This restricts actin nucleation to distal positions within the cell, and in the absence of Multiple Wing Hairs, ectopic actin bundles form across the apical surface (Wong and Adler, 1993). On the distal side, casein kinase 1 γ CK1g/gilamesh is required to further refine prehair nucleation to a single site through a parallel mechanism involving Rab11-dependent vesicle traffic to the site of prehair formation (Gault et al., 2012). Rho and Rho kinase (Drok) have also been implicated in wing hair formation, but their roles are difficult to dissect due to the numerous functions of Rho in cell shape and cell division (Winter et al., 2001; Yan et al., 2009).

Actomyosin contraction and convergent extension (CE).

CE was the first vertebrate process to be linked molecularly to PCP (Wallingford et al., 2000). During CE, mesenchymal cells elongate, form mediolateral-directed protrusions, and intercalate mediolaterally, narrowing the mediolateral axis while simultaneously lengthening the anterior–posterior (A-P) axis (Fig. 3 A; Keller, 2002). Mediolateral polarization, elongation, and intercalation are lost when core PCP components are disrupted, leading to a failure in CE (Tada and Smith, 2000; Wallingford et al., 2000; Goto and Keller, 2002; Jessen et al., 2002). While several PCP-dependent mechanisms have been proposed to mediate CE movements, two recent studies provide direct mechanistic links between asymmetrically localized core PCP components and CE behaviors. In neuroepithelial cells, PCP specifies the localization of myosin to the A-P faces of intercalating cells. Fmi/Celsr1 and Dvl recruit the formin DAAM1 to the A-P junction, which in turn binds and activates PDZ-RhoGEF. This likely activates RhoA and myosin contractility specifically at A-P junctions, resulting in medial-directed cell intercalation and neural plate bending (Nishimura et al., 2012). A similar mechanism was found to drive CE movements of mesenchymal cells during Xenopus gastrulation. In this case, Fritz and Dsh help to localize septins to mediolateral vertices, where they spatially restrict cortical actomyosin contractility and junctional shrinking to A-P cell edges, thus driving cell intercalation (Fig. 3 A; Kim et al., 2010; Shindo and Wallingford, 2014). Together these studies show how asymmetric PCP localization produces collectively polarized cell behaviors through spatial modulation of the cytoskeleton.Open in a separate windowFigure 3.Polarized cell behaviors controlled by PCP. (A) PCP drives convergent extension (CE). CE in vertebrates is driven by mediolateral intercalation, which narrows the mediolateral axis while simultaneously lengthening the A-P axis. Mediolateral intercalation is accompanied by cell polarization and elongation and the formation of mediolateral protrusions, all of which require core PCP function. Pk localizes anteriorly (Ciruna et. al., 2006; Yin et al., 2008), whereas Dsh localizes posteriorly (Yin et. al., 2008). In addition, PCP proteins recruit myosin to A-P cell borders, leading to actomyosin contractility and junctional shrinking. (B) Asymmetric cell division. Drosophila sensory organ precursors (SOPs) divide asymmetrically along the epithelial plane, giving rise to distinct anterior and posterior daughters. Spindle alignment along the A-P axis is PCP dependent. Dsh interacts with Mud/NuMA and the dynein complex posteriorly while Vang links Pins/LGN-Mud/NuMA-dynein on the anterior. This links astral MTs to the A-P cortex, bringing the spindle into register with the A-P axis. (C) Positioning of the kinocilium in the inner ear. The placement of kinocilium in sensory hair cells of the inner ear determines the position of V-shaped stereocilia bundles. Gαi and mPins/LGN localize on the abneural side on the hair cell, where they are required for abneural positioning the MT-based kinocilium. The collective alignment of kinocilia and stereocilia bundles across the epithelium requires the core PCP component Vangl2. Vangl2 (light green) localizes to the abneural side of supporting cells. Whether Fz (dark blue) associates on the opposite face is not yet clear (Ezan et. al., 2013).

Positioning of centrosomes and cilia.

PCP regulates the positioning of MT-based structures including the mitotic spindle and cilia. In Drosophila sensory organ precursors and early zebrafish embryos, PCP controls mitotic spindle orientation along the epithelial plane by interacting with the highly conserved spindle orientation complex, which links astral MTs to the cell cortex through Mud/NuMA-mediated recruitment of the dynein complex (Ségalen et al., 2010). To orient the spindle, posteriorly localized Dsh binds to Mud/NuMA, which recruits the dynein complex and astral MTs to the posterior cortex. On the anterior side, Pins/LGN recruits Mud/NuMA, bringing the spindle into A-P alignment (Fig. 3 B). Similarly, PCP was recently shown to interact with the spindle orientation machinery to position the kinocilium in nondividing cells of the inner ear (Ezan et al., 2013; Tarchini et al., 2013). In vestibular hair cells, Gαi and mPins/LGN localize to the abneural cortex, opposite Vangl2, where they are required for kinocilium positioning and subsequent alignment of stereocilia bundles (Fig. 3 C; Ezan et al., 2013). MT plus ends and dynein also show an abneural bias suggesting that Gαi-mPins/LGN induces pulling on MTs by a similar mechanism that orients the centrosome during spindle orientation. Vangl2 is required for the alignment of Gαi-Pins/LGN crescents between cells, coordinating kinocilia positioning and stereocilia polarity across the tissue (Fig. 3 C; Ezan et al., 2013). Thus the PCP pathway co-opts the spindle orientation machinery to specify not only the division plane but also cilia position in nondividing cells. As PCP is required for asymmetric cilia positioning in a wide range of cell types, including the node (Antic et al., 2010; Borovina et. al., 2010; Song et al., 2010; Hashimoto et. al., 2010), it will be interesting to determine whether this mechanism is conserved.

Concluding remarks

PCP is a fundamental and highly conserved process coordinating a vast number of polarized cell behaviors. While the number of functions ascribed to PCP continues to grow, an understanding of the mechanisms establishing PCP is still far from complete. The development of cellular asymmetry from uniform distributions is not well understood, and will benefit from recent advances in high-resolution, time-lapse imaging with photoconvertible fluorescent tags. Other important issues to resolve include deciphering the structural domains and biochemical interactions mediating intercellular communication, identifying the global cues that orient PCP especially in vertebrates, and deciphering the mechanisms by which complex multicellular structures, like lung branches and hair follicles, are oriented by the PCP machinery.     

The tubulin code: Molecular components,readout mechanisms,and functions

Microtubules are cytoskeletal filaments that are dynamically assembled from α/β-tubulin heterodimers. The primary sequence and structure of the tubulin proteins and, consequently, the properties and architecture of microtubules are highly conserved in eukaryotes. Despite this conservation, tubulin is subject to heterogeneity that is generated in two ways: by the expression of different tubulin isotypes and by posttranslational modifications (PTMs). Identifying the mechanisms that generate and control tubulin heterogeneity and how this heterogeneity affects microtubule function are long-standing goals in the field. Recent work on tubulin PTMs has shed light on how these modifications could contribute to a “tubulin code” that coordinates the complex functions of microtubules in cells.

Introduction

Microtubules are key elements of the eukaryotic cytoskeleton that dynamically assemble from heterodimers of α- and β-tubulin. The structure of microtubules, as well as the protein sequences of α- and β-tubulin, is highly conserved in evolution, and consequently, microtubules look alike in almost all species. Despite the high level of conservation, microtubules adapt to a large variety of cellular functions. This adaptation can be mediated by a large panel of microtubule-associated proteins (MAPs), including molecular motors, as well as by mechanisms that directly modify the microtubules, thus either changing their biophysical properties or attracting subsets of MAPs that convey specific functions to the modified microtubules. Two different mechanism can generate microtubule diversity: the expression of different α- and β-tubulin genes, referred to as tubulin isotypes, and the generation of posttranslational modifications (PTMs) on α- and β-tubulin (Figs. 1 and ​and2).2). Although known for several decades, deciphering how tubulin heterogeneity controls microtubule functions is still largely unchartered. This review summarizes the current advances in the field and discusses new concepts arising.Open in a separate windowFigure 1.Tubulin heterogeneity generated by PTMs. (A) Schematic representation of the distribution of different PTMs of tubulin on the α/β-tubulin dimer with respect to their position in the microtubule lattice. Acetylation (Ac), phosphorylation (P), and polyamination (Am) are found within the tubulin bodies that assemble into the microtubule lattice, whereas polyglutamylation, polyglycylation, detyrosination, and C-terminal deglutamylation take place within the C-terminal tubulin tails that project away from the lattice surface. The tubulin dimer represents TubA1A and TubB2B (Fig. 2), and modification sites for polyglutamylation and polyglycylation have been randomly chosen. (B) Chemical structure of the branched peptide formed by polyglutamylation and polyglycylation, using the γ-carboxyl groups of the modified glutamate residues as acceptor sites for the isopeptide bonds. Note that in the case of polyglutamylation, the elongation of the side chains generates classical peptide bonds (Redeker et al., 1991).Open in a separate windowFigure 2.Heterogeneity of C-terminal tails of tubulin isotypes and their PTMs. The amino acid sequences of all tubulin genes found in the human genome are indicated, starting at the last amino acid of the folded tubulin bodies. Amino acids are represented in single-letter codes and color coded according to their biochemical properties. Known sites for polyglutamylation are indicated (Eddé et al., 1990; Alexander et al., 1991; Rüdiger et al., 1992). Potential modification sites (all glutamate residues) are indicated. Known C-terminal truncation reactions of α/β-tubulin (tub) are indicated. The C-terminal tails of the yeast Saccharomyces cerevisiae are shown to illustrate the phylogenetic diversity of these domains.

Tubulin isotypes

The cloning of the first tubulin genes in the late 1970’s (Cleveland et al., 1978) revealed the existence of multiple genes coding for α- or β-tubulin (Ludueña and Banerjee, 2008) that generate subtle differences in their amino acid sequences, particularly in the C-terminal tails (Fig. 2). It was assumed that tubulin isotypes, as they were named, assemble into discrete microtubule species that carry out unique functions. This conclusion was reinforced by the observation that some isotypes are specifically expressed in specialized cells and tissues and that isotype expression changes during development (Lewis et al., 1985; Denoulet et al., 1986). These high expectations were mitigated by a subsequent study showing that all tubulin isotypes freely copolymerize into heterogeneous microtubules (Lewis et al., 1987). To date, only highly specialized microtubules, such as ciliary axonemes (Renthal et al., 1993; Raff et al., 2008), neuronal microtubules (Denoulet et al., 1986; Joshi and Cleveland, 1989), and microtubules of the marginal band of platelets (Wang et al., 1986; Schwer et al., 2001) are known to depend on some specific (β) tubulin isotypes, whereas the function of most other microtubules appears to be independent of their isotype composition.More recently, a large number of mutations in single tubulin isotypes have been linked to deleterious neurodevelopmental disorders (Keays et al., 2007; Fallet-Bianco et al., 2008; Tischfield et al., 2010; Cederquist et al., 2012; Niwa et al., 2013). Mutations of a single tubulin isotype could lead to an imbalance in the levels of tubulins as a result of a lack of incorporation of mutant isoforms into the microtubule lattice or to incorporation that perturbs the architecture or dynamics of the microtubules. The analysis of tubulin disease mutations is starting to reveal how subtle alterations of the microtubule cytoskeleton can lead to functional aberrations in cells and organisms and might provide novel insights into the roles of tubulin isotypes that have so far been considered redundant.

Tubulin PTMs

Tubulin is subject to a large range of PTMs (Fig. 1), from well-known ones, such as acetylation or phosphorylation, to others that have so far mostly been found on tubulin. Detyrosination/tyrosination, polyglutamylation, and polyglycylation, for instance, might have evolved to specifically regulate tubulin and microtubule functions, in particular in cilia and flagella, as their evolution is closely linked to these organelles. The strong link between those modifications and tubulin evolution has led to the perception that they are tubulin PTMs; however, apart from detyrosination/tyrosination, most of them have other substrates (Regnard et al., 2000; Xie et al., 2007; van Dijk et al., 2008; Rogowski et al., 2009).

Tubulin acetylation.

Tubulin acetylation was discovered on lysine 40 (K40; Fig. 1 A) of flagellar α-tubulin in Chlamydomonas reinhardtii (L’Hernault and Rosenbaum, 1985) and is generally enriched on stable microtubules in cells. Considering that K40 acetylation per se has no effect on the ultrastructure of microtubules (Howes et al., 2014), it is rather unlikely that it directly stabilizes microtubules. As a result of its localization at the inner face of microtubules (Soppina et al., 2012), K40 acetylation might rather affect the binding of microtubule inner proteins, a poorly characterized family of proteins (Nicastro et al., 2011; Linck et al., 2014). Functional experiments in cells have further suggested that K40 acetylation regulates intracellular transport by regulating the traffic of kinesin motors (Reed et al., 2006; Dompierre et al., 2007). These observations could so far not be confirmed by biophysical measurements in vitro (Walter et al., 2012; Kaul et al., 2014), suggesting that in cells, K40 acetylation might affect intracellular traffic by indirect mechanisms.Enzymes involved in K40 acetylation are HDAC6 (histone deacetylase family member 6; Hubbert et al., 2002) and Sirt2 (sirtuin type 2; North et al., 2003). Initial functional studies used overexpression, depletion, or chemical inhibition of these enzymes. These studies should be discussed with care, as both HDAC6 and Sirt2 deacetylate other substrates and have deacetylase-independent functions and chemical inhibition of HDAC6 is not entirely selective for this enzyme (Valenzuela-Fernández et al., 2008). In contrast, acetyl transferase α-Tat1 (or Mec-17; Akella et al., 2010; Shida et al., 2010) specifically acetylates α-tubulin K40 (Fig. 3), thus providing a more specific tool to investigate the functions of K40 acetylation. Knockout mice of α-Tat1 are completely void of K40-acetylated tubulin; however, they show only slight phenotypic aberrations, for instance, in their sperm flagellum (Kalebic et al., 2013). A more detailed analysis of α-Tat1 knockout mice demonstrated that absence of K40 acetylation leads to reduced contact inhibition in proliferating cells (Aguilar et al., 2014). In migrating cells, α-Tat1 is targeted to microtubules at the leading edge by clathrin-coated pits, resulting in locally restricted acetylation of those microtubules (Montagnac et al., 2013). A recent structural study of α-Tat1 demonstrated that the low catalytic rate of this enzyme, together with its localization inside the microtubules, caused acetylation to accumulate selectively in stable, long-lived microtubules (Szyk et al., 2014), thus explaining the link between this PTM and stable microtubules in cells. However, the direct cellular function of K40 acetylation on microtubules is still unclear.Open in a separate windowFigure 3.Enzymes involved in PTM of tubulin. Schematic representation of known enzymes (mammalian enzymes are shown) involved in the generation and removal of PTMs shown in Fig. 1. Note that some enzymes still remain unknown, and some modifications are irreversible. (*CCP5 preferentially removes branching points [Rogowski et al., 2010]; however, the enzyme can also hydrolyze linear glutamate chains [Berezniuk et al., 2013]).Recent discoveries have brought up the possibility that tubulin could be subject to multiple acetylation events. A whole-acetylome study identified >10 novel sites on α- and β-tubulin (Choudhary et al., 2009); however, none of these sites have been confirmed. Another acetylation event has been described at lysine 252 (K252) of β-tubulin. This modification is catalyzed by the acetyltransferase San (Fig. 3) and might regulate the assembly efficiency of microtubules as a result of its localization at the polymerization interface (Chu et al., 2011).

Tubulin detyrosination.

Most α-tubulin genes in different species encode a C-terminal tyrosine residue (Fig. 2; Valenzuela et al., 1981). This tyrosine can be enzymatically removed (Hallak et al., 1977) and religated (Fig. 3; Arce et al., 1975). Mapping of tyrosinated and detyrosinated microtubules in cells using specific antibodies (Gundersen et al., 1984; Geuens et al., 1986; Cambray-Deakin and Burgoyne, 1987a) revealed that subsets of interphase and mitotic spindle microtubules are detyrosinated (Gundersen and Bulinski, 1986). As detyrosination was mostly found on stable and long-lived microtubules, especially in neurons (Cambray-Deakin and Burgoyne, 1987b; Robson and Burgoyne, 1989; Brown et al., 1993), it was assumed that this modification promotes microtubule stability (Gundersen et al., 1987; Sherwin et al., 1987). Although a direct stabilization of the microtubule lattice was considered unlikely (Khawaja et al., 1988), it was found more recently that detyrosination protects cellular microtubules from the depolymerizing activity of kinesin-13–type motor proteins, such as KIF2 or MCAK, thus increasing their longevity (Peris et al., 2009; Sirajuddin et al., 2014).Besides kinesin-13 motors, plus end–tracking proteins with cytoskeleton-associated protein glycine-rich (CAP-Gly) domains, such as CLIP170 or p150/glued, specifically interact with tyrosinated microtubules (Peris et al., 2006; Bieling et al., 2008) via this domain (Honnappa et al., 2006). In contrast, kinesin-1 moves preferentially on detyrosinated microtubules tracks in cells (Liao and Gundersen, 1998; Kreitzer et al., 1999; Konishi and Setou, 2009). The effect of detyrosination on kinesin-1 motor behavior was recently measured in vitro, and a small but significant increase in the landing rate and processivity of the motor has been found (Kaul et al., 2014). Such subtle changes in the motor behavior could, in conjunction with other factors, such as regulatory MAPs associated with cargo transport complexes (Barlan et al., 2013), lead to a preferential use of detyrosinated microtubules by kinesin-1 in cells.Despite the early biochemical characterization of a detyrosinating activity, the carboxypeptidase catalyzing detyrosination of α-tubulin has yet to be identified (Hallak et al., 1977; Argaraña et al., 1978, 1980). In contrast, the reverse enzyme, tubulin tyrosine ligase (TTL; Fig. 3; Raybin and Flavin, 1975; Deanin and Gordon, 1976; Argaraña et al., 1980), has been purified (Schröder et al., 1985) and cloned (Ersfeld et al., 1993). TTL modifies nonpolymerized tubulin dimers exclusively. This selectivity is determined by the binding interface between the TTL and tubulin dimers (Szyk et al., 2011, 2013; Prota et al., 2013). In contrast, the so far unidentified detyrosinase acts preferentially on polymerized microtubules (Kumar and Flavin, 1981; Arce and Barra, 1983), thus modifying a select population of microtubules within cells (Gundersen et al., 1987).In most organisms, only one unique gene for TTL exists. Consequently, TTL knockout mice show a huge accumulation of detyrosinated and particularly Δ2-tubulin (see next section). TTL knockout mice die before birth (Erck et al., 2005) with major developmental defects in the nervous system that might be related to aberrant neuronal differentiation (Marcos et al., 2009). TTL is strictly tubulin specific (Prota et al., 2013), indicating that all observed defects in TTL knockout mice are directly related to the deregulation of the microtubule cytoskeleton.

Δ2-tubulin and further C-terminal modification.

A biochemical study of brain tubulin revealed that ∼35% of α-tubulin cannot be retyrosinated (Paturle et al., 1989) because of the lack of the penultimate C-terminal glutamate residue of the primary protein sequence (Fig. 2; Paturle-Lafanechère et al., 1991). This so-called Δ2-tubulin (for two C-terminal amino acids missing) cannot undergo retyrosination as a result of structural constraints within TTL (Prota et al., 2013) and thus is considered an irreversible PTM.Δ2-tubulin accumulates in long-lived microtubules of differentiated neurons, axonemes of cilia and flagella, and also in cellular microtubules that have been artificially stabilized, for instance, with taxol (Paturle-Lafanechère et al., 1994). The generation of Δ2-tubulin requires previous detyrosination of α-tubulin; thus, the levels of this PTM are indirectly regulated by the detyrosination/retyrosination cycle. This mechanistic link is particularly apparent in the TTL knockout mice, which show massive accumulation of Δ2-tubulin in all tested tissues (Erck et al., 2005). Loss of TTL and the subsequent increase of Δ2-tubulin levels were also linked to tumor growth and might contribute to the aggressiveness of the tumors by an as-yet-unknown mechanism (Lafanechère et al., 1998; Mialhe et al., 2001). To date, no specific biochemical role of Δ2-tubulin has been determined; thus, one possibility is that the modification simply locks tubulin in the detyrosinated state.The enzymes responsible for Δ2-tubulin generation are members of a family of cytosolic carboxypeptidases (CCPs; Fig. 3; Kalinina et al., 2007; Rodriguez de la Vega et al., 2007), and most of them also remove polyglutamylation from tubulin (see next section; Rogowski et al., 2010). These enzymes are also able to generate Δ3-tubulin (Fig. 1 A; Berezniuk et al., 2012), indicating that further degradation of the tubulin C-terminal tails are possible; however, the functional significance of this event is unknown.

Polyglutamylation.

Polyglutamylation is a PTM that occurs when secondary glutamate side chains are formed on γ-carboxyl groups of glutamate residues in a protein (Fig. 1, A and B). The modification was first discovered on α- and β-tubulin from the brain (Eddé et al., 1990; Alexander et al., 1991; Rüdiger et al., 1992; Mary et al., 1994) as well as on axonemal tubulin from different species (Mary et al., 1996, 1997); however, it is not restricted to tubulin (Regnard et al., 2000; van Dijk et al., 2008). Using a glutamylation-specific antibody, GT335 (Wolff et al., 1992), it was observed that tubulin glutamylation increases during neuronal differentiation (Audebert et al., 1993, 1994) and that axonemes of cilia and flagella (Fouquet et al., 1994), as well as centrioles of mammalian centrosomes (Bobinnec et al., 1998), are extensively glutamylated.Enzymes catalyzing polyglutamylation belong to the TTL-like (TTLL) family (Regnard et al., 2003; Janke et al., 2005). In mammals, nine glutamylases exist, each of them showing intrinsic preferences for modifying either α- or β-tubulin as well as for initiating or elongating glutamate chains (Fig. 3; van Dijk et al., 2007). Two of the six well-characterized TTLL glutamylases also modify nontubulin substrates (van Dijk et al., 2008).Knockout or depletion of glutamylating enzymes in different model organisms revealed an evolutionarily conserved role of glutamylation in cilia and flagella. In motile cilia, glutamylation regulates beating behavior (Janke et al., 2005; Pathak et al., 2007; Ikegami et al., 2010) via the regulation of flagellar dynein motors (Kubo et al., 2010; Suryavanshi et al., 2010). Despite the expression of multiple glutamylases in ciliated cells and tissues, depletion or knockout of single enzymes often lead to ciliary defects, particularly in motile cilia (Ikegami et al., 2010; Vogel et al., 2010; Bosch Grau et al., 2013; Lee et al., 2013), suggesting essential and nonredundant regulatory functions of these enzymes in cilia.Despite the enrichment of polyglutamylation in neuronal microtubules (Audebert et al., 1993, 1994), knockout of TTLL1, the major polyglutamylase in brain (Janke et al., 2005), did not show obvious neuronal defects in mice (Ikegami et al., 2010; Vogel et al., 2010). This suggests a tolerance of neuronal microtubules to variations in polyglutamylation.Deglutamylases, the enzymes that reverse polyglutamylation, were identified within a novel family of CCPs (Kimura et al., 2010; Rogowski et al., 2010). So far, three out of six mammalian CCPs have been shown to cleave C-terminal glutamate residues, thus catalyzing both the reversal of polyglutamylation and the removal of gene-encoded glutamates from the C termini of proteins (Fig. 3). The hydrolysis of gene-encoded glutamate residues is not restricted to tubulin, in which it generates Δ2- and Δ3-tubulin, but has also been reported for other proteins such as myosin light chain kinase (Rusconi et al., 1997; Rogowski et al., 2010). One enzyme of the CCP family, CCP5, preferentially removes branching points generated by glutamylation, thus allowing the complete reversal of the polyglutamylation modification (Kimura et al., 2010; Rogowski et al., 2010). However, CCP5 can also hydrolyze C-terminal glutamate residues from linear peptide chains similar to other members of the CCP family (Berezniuk et al., 2013).CCP1 is mutated in a well-established mouse model for neurodegeneration, the pcd (Purkinje cell degeneration) mouse (Mullen et al., 1976; Greer and Shepherd, 1982; Fernandez-Gonzalez et al., 2002). The absence of a key deglutamylase leads to strong hyperglutamylation in brain regions that undergo degeneration, such as the cerebellum and the olfactory bulb (Rogowski et al., 2010). When glutamylation levels were rebalanced by depletion or knockout of the major brain polyglutamylase TTLL1 (Rogowski et al., 2010; Berezniuk et al., 2012), Purkinje cells survived. Although the molecular mechanisms of hyperglutamylation-induced degeneration remain to be elucidated, perturbation of neuronal transport, as well as changes in the dynamics and stability of microtubules, is expected to be induced by hyperglutamylation. Increased polyglutamylation levels have been shown to affect kinesin-1–mediated transport in cultured neurons (Maas et al., 2009), and the turnover of microtubules can also be regulated by polyglutamylation via the activation of microtubule-severing enzymes such as spastin (Lacroix et al., 2010).Subtle differences in polyglutamylation can be seen on diverse microtubules in different cell types. The functions of these modifications remain to be studied; however, its wide distribution strengthens the idea that it could be involved in fine-tuning a range of microtubule functions.

Polyglycylation.

Tubulin polyglycylation or glycylation, like polyglutamylation, generates side chains of glycine residues within the C-terminal tails of α- and β-tubulin (Fig. 1, A and B). The modification sites of glycylation are considered to be principally the same as for glutamylation, and indeed, both PTMs have been shown to be interdependent in cells (Rogowski et al., 2009; Wloga et al., 2009). Initially discovered on Paramecium tetraurelia tubulin (Redeker et al., 1994), glycylation has been extensively studied using two antibodies, TAP952 and AXO49 (Bressac et al., 1995; Levilliers et al., 1995; Bré et al., 1996). In contrast to polyglutamylation, glycylation is restricted to cilia and flagella in most organisms analyzed so far.Glycylating enzymes are also members of the TTLL family, and homologues of these enzymes have so far been found in all organisms with proven glycylation of ciliary axonemes (Rogowski et al., 2009; Wloga et al., 2009). In mammals, initiating (TTLL3 and TTLL8) and elongating (TTLL10) glycylases work together to generate polyglycylation (Fig. 3). In contrast, the two TTLL3 orthologues from Drosophila melanogaster can both initiate and elongate glycine side chains (Rogowski et al., 2009).In mice, motile ependymal cilia in brain ventricles acquire monoglycylation upon maturation, whereas polyglycylation is observed only after several weeks (Bosch Grau et al., 2013). Sperm flagella, in contrast, acquire long glycine chains much faster, suggesting that the extent of polyglycylation could correlate with the length of the axonemes (Rogowski et al., 2009). Depletion of glycylases in mice (ependymal cilia; Bosch Grau et al., 2013), zebrafish (Wloga et al., 2009; Pathak et al., 2011), Tetrahymena thermophila (Wloga et al., 2009), and D. melanogaster (Rogowski et al., 2009) consistently led to ciliary disassembly or severe ciliary defects. How glycylation regulates microtubule functions remains unknown; however, the observation that glycylation-depleted axonemes disassemble after initial assembly (Rogowski et al., 2009; Bosch Grau et al., 2013) suggests a role of this PTM in stabilizing axonemal microtubules. Strikingly, human TTLL10 is enzymatically inactive; thus, humans have lost the ability to elongate glycine side chains (Rogowski et al., 2009). This suggests that the elongation of the glycine side chains is not an essential aspect of the function of this otherwise critical tubulin PTM.

Other tubulin PTMs.

Several other PTMs have been found on tubulin. Early studies identified tubulin phosphorylation (Eipper, 1974; Gard and Kirschner, 1985; Díaz-Nido et al., 1990); however, no specific functions were found. The perhaps best-studied phosphorylation event on tubulin takes place at serine S172 of β-tubulin (Fig. 1 A), is catalyzed by the Cdk1 (Fig. 3), and might regulate microtubule dynamics during cell division (Fourest-Lieuvin et al., 2006; Caudron et al., 2010). Tubulin can be also modified by the spleen tyrosine kinase Syk (Fig. 3; Peters et al., 1996), which might play a role in immune cells (Faruki et al., 2000; Sulimenko et al., 2006) and cell division (Zyss et al., 2005; Sulimenko et al., 2006).Polyamination has recently been discovered on brain tubulin (Song et al., 2013), after having been overlooked for many years as a result of the low solubility of polyaminated tubulin. Among several glutamine residues of α- and β-tubulin that can be polyaminated, Q15 of β-tubulin is considered the primary modification site (Fig. 1 A). Polyamination is catalyzed by transglutaminases (Fig. 3), which modify free tubulin as well as microtubules in an irreversible manner, and most likely contribute to the stabilization of microtubules (Song et al., 2013).Tubulin was also reported to be palmitoylated (Caron, 1997; Ozols and Caron, 1997; Caron et al., 2001), ubiquitinated (Ren et al., 2003; Huang et al., 2009; Xu et al., 2010), glycosylated (Walgren et al., 2003; Ji et al., 2011), arginylated (Wong et al., 2007), methylated (Xiao et al., 2010), and sumoylated (Rosas-Acosta et al., 2005). These PTMs have mostly been reported without follow-up studies, and some of them are only found in specific cell types or organisms and/or under specific metabolic conditions. Further studies will be necessary to gain insights into their potential roles for the regulation of the microtubule cytoskeleton.

Current advances and future perspectives

The molecular heterogeneity of microtubules, generated by the expression of different tubulin isotypes and by the PTM of tubulin has fascinated the scientific community for ∼40 years. Although many important advances have been made in the past decade, the dissection of the molecular mechanisms and a comprehensive understanding of the biological functions of tubulin isotypes and PTMs will be a challenging field of research in the near future.

Direct measurements of the impact of tubulin heterogeneity.

The most direct and reliable type of experiments to determine the impact of tubulin heterogeneity on microtubule behavior are in vitro measurements with purified proteins. However, most biophysical work on microtubules has been performed with tubulin purified from bovine, ovine, or porcine brains, which can be obtained in large quantities and with a high degree of purity and activity (Vallee, 1986; Castoldi and Popov, 2003). Brain tubulin is a mixture of different tubulin isotypes and is heavily posttranslationally modified and thus inept for investigating the functions of tubulin heterogeneity (Denoulet et al., 1986; Cambray-Deakin and Burgoyne, 1987b; Paturle et al., 1989; Eddé et al., 1990). Thus, pure, recombinant tubulin will be essential to dissect the roles of different tubulin isoforms and PTMs.Attempts to produce recombinant, functional α- and β-tubulin in bacteria have failed so far (Yaffe et al., 1988), most likely because of the absence of the extensive tubulin-specific folding machinery (Yaffe et al., 1992; Gao et al., 1993; Tian et al., 1996; Vainberg et al., 1998) in prokaryotes. An alternative source of tubulin with less isotype heterogeneity and with almost no PTMs is endogenous tubulin from cell lines such as HeLa, which in the past has been purified using a range of biochemical procedures (Bulinski and Borisy, 1979; Weatherbee et al., 1980; Farrell, 1982; Newton et al., 2002; Fourest-Lieuvin, 2006). Such tubulin can be further modified with tubulin-modifying enzymes, such as polyglutamylases, either by expressing those enzymes in the cells before tubulin purification (Lacroix and Janke, 2011) or in vitro with purified enzymes (Vemu et al., 2014). Despite some technical limitations of these methods, HeLa tubulin modified in cells has been successfully used in an in vitro study on the role of polyglutamylation in microtubule severing (Lacroix et al., 2010).Naturally occurring variants of tubulin isotypes and PTMs can be purified from different organisms, organs, or cell types, but obviously, only some combinations of tubulin isotypes and PTMs can be obtained by this approach. The recent development of an affinity purification method using the microtubule-binding TOG (tumor overexpressed gene) domain of yeast Stu2p has brought a new twist to this approach, as it allows purifying small amounts of tubulin from any cell type or tissue (Widlund et al., 2012).The absence of tubulin heterogeneity in yeast has made budding and fission yeast potential expression systems for recombinant, PTM-free tubulin (Katsuki et al., 2009; Drummond et al., 2011; Johnson et al., 2011). However, the expression of mammalian tubulin in this system has remained impossible. This problem was then partially circumvented by expressing tubulin chimeras that consist of a yeast tubulin body fused to mammalian C-terminal tubulin tails, thus mimicking different tubulin isotypes (Sirajuddin et al., 2014). Moreover, detyrosination can be generated by deleting the key C-terminal residue from endogenous or chimeric α-tubulin (Badin-Larçon et al., 2004), and polyglutamylation is generated by chemically coupling glutamate side chains to specifically engineered tubulin chimeras (Sirajuddin et al., 2014). These approaches allowed the first direct measurements of the impact of tubulin isotypes and PTMs on the behavior of molecular motors in vitro (Sirajuddin et al., 2014) and the analysis of the effects of tubulin heterogeneity on microtubule behavior and interactions inside the yeast cell (Badin-Larçon et al., 2004; Aiken et al., 2014).Currently, the most promising development has been the successful purification of fully functional recombinant tubulin from the baculovirus expression system (Minoura et al., 2013). Using this system, defined α/β-tubulin dimers can be obtained using two different epitope tags on α- and β-tubulin, respectively. Although these epitope tags are essential for separating recombinant from the endogenous tubulin, they could also affect tubulin assembly or microtubule–MAP interactions. Thus, future developments should focus on eliminating these tags.Current efforts have brought the possibility of producing recombinant tubulin into reach. Further improvement and standardization of these methods will certainly provide a breakthrough in understanding the mechanisms by which tubulin heterogeneity contributes to microtubule functions.

Complexity of tubulin—understanding the regulatory principles.

The diversity of tubulin genes (isotypes) and the complexity of tubulin PTMs have led to the proposal of the term “tubulin code” (Verhey and Gaertig, 2007; Wehenkel and Janke, 2014), in analogy to the previously coined histone code (Jenuwein and Allis, 2001). Tubulin molecules consist of a highly structured and thus evolutionarily conserved tubulin body and the unstructured and less conserved C-terminal tails (Nogales et al., 1998). As PTMs and sequence variations within the tubulin body are expected to affect the conserved tubulin fold and therefore the properties of the microtubule lattice, they are not likely to be involved in generating the tubulin code. In contrast, modulations of the C-terminal tails could encode signals on the microtubule surface without perturbing basic microtubule functions and properties (Figs. 1 A and ​and4).4). Indeed, the highest degree of gene-encoded diversity (Fig. 2) and the highest density and complexity of PTMs (Fig. 1) are found within these tail domains.Open in a separate windowFigure 4.Molecular components of the tubulin code. Schematic representation of potential coding elements that could generate specific signals for the tubulin code. (A) The length of the C-terminal tails of different tubulin isotypes differ significantly (Fig. 2) and could have an impact on the interactions between microtubules and MAPs. (B) Tubulin C-terminal tails are rich in charged amino acid residues. The distribution of these residues and local densities of charges could influence the electrostatic interactions with the tails and the readers. (C) Although each glutamate residue within the C-terminal tails could be considered a potential modification site, only some sites have been found highly occupied in tubulin purifications from native sources. This indicates selectivity of the modification reactions, which can participate in the generation of specific modification patterns (see D). Modification sites might be distinguished by their neighboring amino acid residues, which could create specific modification epitopes. (D) As a result of the large number of modification sites and the variability of side chains, a large variety of modification patterns could be generated within a single C-terminal tail of tubulin. (E) Modification patterns as shown in D can be distinct between α- and β-tubulin. These modification patterns could be differentially distributed at the surface of the microtubule lattice, thus generating a higher-order patterning. Tub, tubulin. For color coding, see Fig. 2.Considering the number of tubulin isotypes plus all potential combinations of PTMs (e.g., each glutamate residue within the C-terminal tubulin tail could be modified by either polyglutamylation or polyglycylation, each of them generating side chains of different lengths; Fig. 4), the number of distinct signals generated by the potential tubulin code would be huge. However, as many of these potential signals represent chemical structures that are similar and might not be reliably distinguished by readout mechanisms, it is possible that the tubulin code generates probabilistic signals. In this scenario, biochemically similar modifications would have similar functional readouts, and marginal differences between those signals would only bias biological processes but not determine them. This stands in contrast to the concept of the histone code, in which precise patterns of different PTMs on the histone proteins encode distinct biological signals.The concept of probabilistic signaling is already inscribed in the machinery that generates the tubulin code. Polyglutamylases and polyglycylases from the TTLL family have preferential activities for either α- or β-tubulin and for generating different lengths of the branched glutamate or glycine chains. Although under conditions of low enzyme concentrations, as found in most cells and tissues, the enzymes seem to selectively generate their preferential type of PTM, higher enzyme concentrations induce a more promiscuous behavior, leading, for instance, to a loss of selectivity for α- or β-tubulin (van Dijk et al., 2007). Similarly, the modifying enzymes might prefer certain modification sites within the C-terminal tails of tubulin but might be equally able to modify other sites, which could be locally regulated in cells. For example, β-tubulin isotypes isolated from mammalian brain were initially found to be glutamylated on single residues (Alexander et al., 1991; Rüdiger et al., 1992), which in the light of the comparably low sensitivity of mass spectrometry at the time might rather indicate a preferential than a unique modification of these sites. Nevertheless, the neuron-specific polyglutamylase for β-tubulin TTLL7 (Ikegami et al., 2006) can incorporate glutamate onto many more modification sites of β-tubulin in vitro (Mukai et al., 2009), which clearly indicates that not all of the possible modification events take place under physiological conditions.Several examples supporting a probabilistic signaling mode of the tubulin code are found in the recent literature. In T. thermophila, a ciliate without tubulin isotype diversity (Gaertig et al., 1993) but with a huge repertoire of tubulin PTMs and tubulin-modifying enzymes (Janke et al., 2005), tubulin can be easily mutagenized to experimentally eliminate sites for PTMs. Mutagenesis of the most commonly occupied glutamylation/glycylation sites within the β-tubulin tails did not generate a clear decrease of glycylation levels nor did it cause obvious phenotypic alterations. This indicates that the modifying enzymes can deviate toward alternative modification sites and that similar PTMs on different sites can compensate the functions of the mutated site. However, when all of the key modification sites were mutated, glycylation became prominently decreased, which led to severe phenotypes, including lethality (Xia et al., 2000). Most strikingly, these phenotypes could be recovered by replacing the C-terminal tail of α-tubulin with the nonmutated β-tubulin tail. This α–β-tubulin chimera became overglycylated and functionally compensated for the absence of modification sites on β-tubulin. The conclusion of this study is that PTM- and isotype-generated signals can fulfill a biological function within a certain range of tolerance.But how efficient is such compensation? The answer can be found in a variety of already described deletion mutants for tubulin-modifying enzymes in different model organisms. Most single-gene knockouts for TTLL genes (glutamylases or glycylases) did not result in prominent phenotypic alterations in mice, even for enzymes that are ubiquitously expressed. Only some highly specialized microtubule structures show functional aberrations upon the deletion of a single enzyme. These “tips of the iceberg” are usually the motile cilia and sperm flagella, which carry very high levels of polyglutamylation and polyglycylation (Bré et al., 1996; Kann et al., 1998; Rogowski et al., 2009). It thus appears that some microtubules are essentially dependent on the generation of specific PTM patterns, whereas others can tolerate changes and appear to function normally. How “normal” these functions are remains to be investigated in future studies. It is possible that defects are subtle and thus overlooked but could become functionally important under specific conditions.A tubulin code also requires readout mechanisms. The most likely “readers” of the tubulin code are MAPs and molecular motors. Considering the probabilistic signaling hypothesis, the expected effects of the signals would be in most cases rather gradual changes, for instance, to fine-tune molecular motor traffic and/or to bias motors toward defined microtubule tracks but not to obliterate motor activity or MAP binding to microtubules. An in vitro study using recombinant tubulin chimeras purified from yeast confirmed this notion (Sirajuddin et al., 2014). By analyzing which elements of the tubulin code can regulate the velocity and processivity of the molecular motors kinesin and dynein, these researchers found that the C-terminal tails of α- and β-tubulin differentially influence the kinetic parameters of the tested motors; however, the modulation was rather modest. One of their striking observations was that a single lysine residue, present in the C-terminal tails of two β-tubulin isotypes (Figs. 2 and ​and4),4), significantly affected motor traffic and that this effect can be counterbalanced by polyglutamylation. These observations are the first in vitro evidence for the interdependence of different elements of the tubulin code and provide another indication for its probabilistic mode of signaling.

Future directions.

One of the greatest technological challenges to understanding the function of the tubulin code is to detect and interpret subtle and complex regulatory events generated by this code. It will thus be instrumental to further develop tools to better distinguish graded changes in PTM levels on microtubules in cells and tissues (Magiera and Janke, 2013) and to reliably measure subtle modulations of microtubule behavior in reconstituted systems.The current advances in the field and especially the availability of whole-organism models, as well as first insights into the pathological role of tubulin mutations (Tischfield et al., 2011), are about to transform our way of thinking about the regulation of microtubule cytoskeleton. Tubulin heterogeneity generates complex probabilistic signals that cannot be clearly attributed to single biological functions in most cases and that are not essential for most cellular processes. Nevertheless, it has been conserved throughout evolution of eukaryotes and can hardly be dismissed as not important. To understand the functional implications of these processes, we might be forced to reconsider how we define biologically important events and how we measure events that might encode probabilistic signals. The answers to these questions could provide novel insights into how complex systems, such as cells and organisms, are sustained throughout difficult and challenging life cycles, resist to environmental stress and diseases, and have the flexibility needed to succeed in evolution.     

Cell biology in development: Cell division and the maintenance of epithelial order

Epithelia are polarized layers of adherent cells that are the building blocks for organ and appendage structures throughout animals. To preserve tissue architecture and barrier function during both homeostasis and rapid growth, individual epithelial cells divide in a highly constrained manner. Building on decades of research focused on single cells, recent work is probing the mechanisms by which the dynamic process of mitosis is reconciled with the global maintenance of epithelial order during development. These studies reveal how symmetrically dividing cells both exploit and conform to tissue organization to orient their mitotic spindles during division and establish new adhesive junctions during cytokinesis.The association of large numbers of cells in tightly organized epithelial layers is a unique and defining feature of Metazoa. Although classical studies of development once labeled distinct embryonic regions as territories, fields, layers, placodes, and primordia, we now know many of these structures to be primarily constructed from epithelial sheets. Epithelial structure and function are critically dependent on cell polarization, which is coupled to the targeted assembly of adhesive junctions along the apicolateral membranes of adjacent cells (Tepass et al., 2001; Cavey and Lecuit, 2009). In brief, the plasma membrane of epithelial cells is polarized into apical and basolateral domains, each enriched with distinct lipid and protein components (Fig. 1; Rodriguez-Boulan et al., 2005; St Johnston and Ahringer, 2010). At the molecular level, E-cadherins are the major class of adhesion proteins that establish cell–cell connections through homophilic interaction across cell membranes (Takeichi, 1991, 2011; Halbleib and Nelson, 2006; Harris and Tepass, 2010). Whereas E-cadherin is apically enriched in invertebrate epithelia, it is localized along the lateral domain of vertebrate epithelial cells. In both cases, E-cadherin interacts with cytoplasmic actin filaments via the catenin class of adaptor proteins, thus coupling intercellular adhesive contacts to the cytoskeleton (Cavey and Lecuit, 2009; Harris and Tepass, 2010; Gomez et al., 2011). Within this framework, the maintenance of both polarity and cell–cell adhesion are essential for epithelial barrier function and tissue architecture during growth and morphogenesis (Papusheva and Heisenberg, 2010; Guillot and Lecuit, 2013b).Open in a separate windowFigure 1.Architectural implications of orthogonal and planar spindle orientations during epithelial cell division. (A) Programmed orthogonal orientation of the mitotic spindle can promote epithelial stratification, although the remodeling of adhesion and polarity complexes during this process remains an important area for further study. (B) Planar spindle orientation is coordinated with the overall cell polarity machinery and thus facilitates conservation of monolayer organization during rapid cell proliferation.During development, epithelia expand by the combined effects of cell growth (increase in cell size) and cell division (increase in cell numbers). Division events are typically oriented either parallel or orthogonal to the plane of the layer and less frequently at oblique angles (Gillies and Cabernard, 2011). When cells divide orthogonally (perpendicular to the plane of the epithelium), the two daughters will be at least initially nonequivalent with respect to position within the cell layer (Fig. 1 A). Under normal conditions, such programmed orthogonal divisions can be used to effect asymmetric segregation of cell fates or to establish distinct cell types, such as in the developing cortex (Fietz et al., 2010; Hansen et al., 2010) or during morphogenesis of stratified epithelia (Lechler and Fuchs, 2005; Williams et al., 2011). Conversely, when cells divide parallel to the plane of the epithelium (planar orientation; Fig. 1 B), both daughter cells are equivalent with respect to mother cell polarity and tightly integrated in the growing monolayer (Morin and Bellaïche, 2011).During planar division, epithelial cells typically round up, constrict in the middle to form the cytokinetic furrow, and divide symmetrically with respect to the apicobasal axis to produce two equal daughter cells. These daughters construct new cell–cell junctions at their nascent interface, thus integrating into the monolayer (Fig. 2, A–G). Although the intricate relationship between cell polarity and cell division has been explored for many years in the context of asymmetric cell division (Rhyu and Knoblich, 1995; Siller and Doe, 2009; Williams and Fuchs, 2013), recent studies have also begun to explore how epithelia maintain their morphology, integrity, and barrier function during continuous rounds of planar cell division and junction assembly. In this review, we highlight recent findings that provide new insights into the problem of symmetric planar cell division in diverse polarized epithelia, with a focus on two crucial mitotic events: (1) the orientation of cell division and (2) the formation of new cell junctions.Open in a separate windowFigure 2.Progression of planar cell division in an epithelial monolayer. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing cell (red). (A) At the level of apical junctions, cells are packed in a polygonal cell arrangement during interphase. (B) In prophase, the dividing nucleus begins to translocate apically as the cell rounds up and maintains a thin basal projection enriched with nonmuscle myosin II and actin (light blue). Notably, this type of nuclear migration is typically observed in pseudostratified columnar epithelia and does not occur in cuboidal and squamous epithelial tissues. (C) Localized molecular landmarks (apical complexes marked as gray bars on cell sides) direct orientation of the mitotic spindle to the plane of the epithelium (arrows). (D) Within the plane of the cell layer, the spindle can be further oriented (arrows) in response to molecular cues, global tissue tension, and local cell geometry. (E and F) After chromosome segregation during anaphase, the cell constricts in the middle and cleaves orthogonal to the plane of the monolayer. (G) After cytokinesis, daughter nuclei move basally and daughter cells form new junctions at their nascent interface (white) while elongating along the apicobasal axis.

Mitotic spindle position and orientation in epithelial cells

Planar orientation of epithelial cell division requires coordinated interaction between the cell polarization machinery and the mitotic spindle itself (Morin and Bellaïche, 2011). In animal cells, the spindle is organized by two symmetrically positioned poles or centrosomes, which nucleate three forms of microtubules (Tanaka, 2010): kinetochore microtubules that attach to the chromosomes, polar microtubules that overlap in an antiparallel fashion over the midplane, and astral microtubules that extend to the cell cortex, which is the actin-rich layer beneath the cell membrane (Lancaster and Baum, 2014). Work in Drosophila melanogaster and vertebrates reveals that at least three factors influence the orientation of this spindle machinery with respect to polarized epithelial architecture: cytoskeletal forces, localized cortical cues, and tissue tension.

Cytoskeletal forces position mitotic nuclei near the apical cell membrane.

In columnar and pseudostratified epithelia where cells elongate along their apicobasal axes, mitotic events are typically restricted to the apical domain of the epithelium (corresponding to the apical membrane of each cell; Fig. 2, C–F). How does the mitotic nucleus achieve the correct apical position? In Drosophila wing discs and zebrafish neuroepithelia, mitotic nuclei and the bulk of the cell cytoplasm are driven apically by actomyosin-dependent cortical contractility at prophase entry (Norden et al., 2009; Leung et al., 2011; Meyer et al., 2011). These events are fundamentally similar to mitotic cell rounding in tissue culture cells (Kunda and Baum, 2009; Lancaster et al., 2013; Lancaster and Baum, 2014). In many epithelia, as the cell rounds up and the nucleus translocates apically, a thin actin-rich projection maintains contact with the basal lamina (Fig. 2, B and C). It remains poorly understood how this structure behaves during cleavage and whether this basal process plays any role in the correct reintegration of the postmitotic daughter cells into the monolayer. Although actomyosin may be the primary driver of apical rounding in many cases, evidence also supports a role for microtubule-based mechanisms in the positioning of premitotic nuclei. In chicken neural tube and mouse cerebral cortex, nuclei migrate apically on microtubules before actomyosin-dependent rounding (Spear and Erickson, 2012a). Centrosomes provide directionality to the microtubules on which the nucleus migrates and organize the spindle once the mitotic chromatin reaches the apical domain (Peyre et al., 2011; Spear and Erickson, 2012a; Nakajima et al., 2013). Collectively, current evidence suggests that both actomyosin- and microtubule-dependent forces conspire to effect mitotic nuclear translocation in a highly context- and species-specific manner. One possibility is that the varying physical dimensions of epithelial cells require varying mechanisms for apical nuclear translocation. For example, highly elongated radial glial cells require active transport of the nucleus on microtubules before mitotic rounding, whereas cortical actomyosin contractility may be sufficient in less elongated cells (Spear and Erickson, 2012b). A major outstanding problem is how cortical contractility triggers cell rounding that is polarized along the apicobasal axis of the cell. Whereas centrosomes function as an apical landmark for nuclei moving on microtubules, it remains unclear what provides the directionality for the basal-to-apical actomyosin contraction. One hypothesis is that certain proteins can restrict the localization of nonmuscle myosin II at the basal domain of epithelial cells. The microcephaly protein Asp interacts with myosin II and regulates its polarized localization along the apicobasal axis in the fly optic lobe neuroepithelium. In asp mutant flies, myosin II is enriched apically instead of basally. Many dividing nuclei fail to reach the apical domain and are thus broadly distributed along the apicobasal axis of the epithelium, leading to a disorganized tissue (Rujano et al., 2013). Interestingly, Asp also interacts with microtubules, associates with spindle poles, and is essential for positioning the spindle in fly and vertebrate epithelia (Saunders et al., 1997; do Carmo Avides and Glover, 1999; Wakefield et al., 2001; Fish et al., 2006). Elucidating the function of proteins such as Asp at the interface of microtubules and actomyosin will be essential to our understanding of how the cytoskeleton drives apical mitotic rounding.

Localized molecular landmarks direct planar spindle orientation.

In most animal cells, the mitotic spindle is anchored to the cell cortex by astral microtubules (Fig. 2, C–E; Théry and Bornens, 2006). Translocation of the dynein–dynactin motor toward the astral microtubule minus ends provides a pulling force on centrosomes and is essential for spindle orientation and pole separation during cell division (Dujardin and Vallee, 2002; Kotak et al., 2012). Molecular cues embedded in the cortex can thus determine spindle orientation by anchoring the dynein–dynactin complex in restricted domains. In cultured MDCK and chick neuroepithelia cells, the Gαi–LGN–nuclear mitotic apparatus (NuMA) complex serves this function (Busson et al., 1998; Hao et al., 2010; Zheng et al., 2010; Peyre et al., 2011). Knockdown or mislocalization of these factors leads to spindle orientation defects that ultimately lead to removal of cell progenitors from the monolayer (Peyre et al., 2011). LGN (Pins in Drosophila) localizes to the lateral cell cortex by binding to the membrane-bound Gαi and enforces spindle orientation by recruiting NuMA (Mud in Drosophila), which binds directly to the dynein–dynactin motor. In certain epithelia, including MDCK cells and Drosophila wing discs, LGN is excluded from the apical domain by atypical PKC (aPKC) phosphorylation, thus restricting it at the lateral cell cortex (Konno et al., 2008; Hao et al., 2010; Zheng et al., 2010; Guilgur et al., 2012). In chick neuroepithelia, however, LGN is restricted at the lateral cortex independently of aPKC, suggesting that other cues control its localization (Morin et al., 2007; Peyre et al., 2011). In the mouse embryonic neocortex, the actin–membrane linkers ERM (ezrin/radixin/moesin) promote the association of LGN with NuMA (Machicoane et al., 2014), indicating that organized cortical actin is critical for correct LGN localization.Cell–cell junctions have been implicated in planar cell division in mammalian epithelia, suggesting a possible direct link between the polarity apparatus and the spindle machinery (Reinsch and Karsenti, 1994; den Elzen et al., 2009). Interfering with E-cadherin function or reducing E-cadherin levels abolishes junctional localization of APC (adenomatous polyposis coli), a microtubule-interacting protein that is required for planar spindle orientation and chromosome alignment (Green et al., 2005; den Elzen et al., 2009). However, spindle orientation may not directly depend on E-cadherin or adherens junctions (AJs) in all cases. In Drosophila follicle cells and imaginal discs as well as Xenopus laevis embryonic epithelia, mitotic spindles exhibit planar orientation but do not align with the AJs (Woolner and Papalopulu, 2012; Bergstralh et al., 2013; Nakajima et al., 2013). Moreover, disruption of AJs in Drosophila follicle cells does not affect spindle position (Bergstralh et al., 2013).In Drosophila wing discs, the spindle poles localize in close proximity to septate junctions, which are positioned immediately basal to AJs (Nakajima et al., 2013). Septate junctions are enriched with many proteins, including the neoplastic tumor suppressors SCRIB (Scribbled) and DLG1 (Discs large 1; Bilder and Perrimon, 2000; Bilder et al., 2000). In asymmetrically dividing cells, such as Drosophila sensory organ precursors and neuroblasts, DLG1 interacts with LGN at the cortex and is required for proper spindle orientation (Bellaïche et al., 2001; Siegrist and Doe, 2005; Johnston et al., 2009). Recent findings indicate that DLG1 is also essential for planar spindle orientation in the symmetric division of epithelial cells. In wing discs, knockdown of scrib or dlg1 leads to randomized spindle orientations. scrib knockdown wing discs exhibit diffuse DLG1 localization but no obvious apicobasal polarity defect, suggesting that epithelial disorganization could be a consequence of aberrant spindle orientation (Nakajima et al., 2013). However, it is not clear whether the septate junctions themselves are important. In Drosophila follicle epithelial cells where septate junctions do not form until relatively late in development (Oshima and Fehon, 2011), DLG1 is localized at the lateral cell cortex and is essential for planar spindle orientation (Bergstralh et al., 2013). Interestingly, dlg1 mutant follicle cells display misoriented divisions yet normal epithelial polarity and tissue organization. In this case, planar spindle orientation appears to be independent of junctions per se but still depends on a DLG1–LGN–NuMA complex, similar to asymmetrically dividing cells (Bergstralh et al., 2013).

Global stress and local cell geometry influence mitotic spindle orientation within the plane of the epithelium.

During planar divisions, the mitotic spindle aligns to the plane of the epithelium (xz; Fig. 2 C) and also within the plane of the cell layer (xy; Fig. 2 D). Studies in gastrulating zebrafish embryos revealed a role for the Wnt–Frizzled–planar cell polarity signaling pathway in orienting cell divisions (Concha and Adams, 1998; Gong et al., 2004). Similarly, the atypical cadherins Fat and Dachsous are involved in orienting cell divisions in the Drosophila wing and in developing mouse kidneys (Baena-López et al., 2005; Saburi et al., 2008). Although both of these pathways have been reviewed elsewhere (Morin and Bellaïche, 2011), recent studies also point to at least two other mechanisms that may independently influence spindle orientation within the plane of the monolayer: (1) global tissue stress and (2) local epithelial cell geometry.Epithelial cell shape and spindle orientation are modulated by global stress that accumulates during tissue growth. In Drosophila wing discs, cells in the center of the wing blade primordium proliferate at a faster rate than in the periphery. Consequently, cells in the periphery are mechanically stretched, and cells in the center are compressed. As a result of stretching, peripheral cells localize myosin II at their cortex and align their mitotic spindle with the stretch axis (LeGoff et al., 2013; Mao et al., 2013). Similarly, epithelial cells of the enveloping cell layer in gastrulating zebrafish embryos elongate and orient their spindle along the direction of tension generated by spreading during epiboly (Campinho et al., 2013). It is unclear whether myosin II directly conveys cell tension to the mitotic apparatus, and it will be necessary to dissect whether cell elongation alone or additional mechanosensing pathways signal cell tension to the mitotic spindle. Keratinocytes from the mammalian epidermis reorient their mitotic spindle in response to mechanical stretch in a NuMA-dependent manner. The mitotic spindle aligns with the cortical NuMA-localized crescent upon stretch and fails to orient when NuMA levels are reduced (Seldin et al., 2013). In summary, global tension generated by growth and cell spreading impact division orientation, suggesting that shape changes in proximity to dividing cells may also lead to a similar effect.Although variations certainly exist, the apical surfaces of proliferating epithelia tend to feature a consistent percentage of hexagonal, pentagonal, heptagonal, and octagonal cell shapes (Gibson et al., 2006; Farhadifar et al., 2007; Aegerter-Wilmsen et al., 2010). In Drosophila imaginal discs, these local patterns of cell packing may systematically influence spindle orientation, as mitotic cells are biased toward cleaving their common interfaces with subhexagonal neighbors (less than six sides) and avoid cleaving their interfaces with superhexagonal neighbors (more than six sides; Gibson et al., 2011). Although the mechanisms underlying the effect of local cell geometry remain elusive, cell packing influences mitotic cell shape and the distribution of adhesive cues, both of which could, in turn, bias spindle orientation. Indeed, dividing cells maintain contacts with their neighbors, which can influence the cell cortex and direct spindle orientation (Goldstein, 1995; Wang et al., 1997). The distribution of adhesions between epithelial cells may also alter the position or action of cortical force generators that interact with spindle microtubules in the mitotic cell. In support of this idea, when single cells are placed on micropatterned substrates, they orient their spindle relative to the geometry of their adhesion pattern and not their cell shape (Théry et al., 2005, 2007). Alternatively, neighbors of different polygonal shapes could stretch the mitotic cell, thus imposing a bias on its long axis. Indeed, sea urchin embryos orient their spindles to divide their longest axis (Hertwig, 1884) and can even sense complex cell geometries to orient their spindles accordingly (Minc et al., 2011). Still, precisely how the interphase morphology of epithelial cells might impinge on mitotic spindle orientation remains an open question.

Genesis of nascent junctions during epithelial cell division

After spindle orientation, the essential processes of cytokinesis and abscission are driven by the assembly and contraction of an actomyosin ring positioned in the cleavage plane (Fededa and Gerlich, 2012). In epithelia, ring contraction accompanied by membrane invagination ultimately gives rise to a new junctional interface between nascent daughter cells. Precisely how this new interface forms remains poorly understood. Recent studies in Drosophila epithelia reveal that, during cytokinesis, (a) E-cadherin levels are reduced at the interface between the cleavage furrow of dividing cells and their neighbors (Fig. 3), and (b) neighbor tension and midbody position guide establishment of new AJs in context with local epithelial geometry (Fig. 4).Open in a separate windowFigure 3.Cytokinetic membrane dynamics in epithelial cells. (A) Cytokinesis of a dividing epithelial cell (yellow) presents several unique structural considerations not addressed by the analysis of single cells. Recent studies (Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013, 2014) report a local reduction of E-cadherin levels in proximity to the contractile ring in the dividing cell and its neighbor (red). How cytokinesis is resolved from there may vary in a context-dependent manner. (B) In Drosophila embryos, ring contraction leads to E-cadherin disengagement, and a gap forms between the mitotic cell and its neighbor (Guillot and Lecuit, 2013a). (C) In the Drosophila pupal notum, the contractile ring pulls the neighbor cell plasma membrane into the cleavage furrow, perhaps enabled by uncoupling of the membrane and the cortex in the neighbor (Herszterg et al., 2013, 2014).Open in a separate windowFigure 4.New AJ formation in dividing epithelial cells. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing epithelial cell (red). (A) Opposing forces (black vertical arrows) develop between the contractile ring in the dividing cell and the two neighboring cells (orange) in proximity to the cleavage furrow. E-cadherin clusters are reduced at the furrow/neighbor interface. (B) Myosin II and tension build up in the neighboring cell, causing the nascent daughter cells to juxtapose their plasma membranes at the presumptive site of junction assembly (black horizontal arrows). (C) Arp2/3 and Rac1 drive actin polymerization at the daughter cell interface around the midbody, stabilizing the nascent junction as the neighboring cell membrane withdraws. (D) The new junction is complete and of suitable length in context with the local epithelial geometry.

Mitotic cells remodel their adhesion junctions during cytokinesis.

Two kinds of forces are at work during cytokinesis: an active force in the dividing cell caused by ring contraction and a reactive force in contacting neighbors caused by their resistance to pulling to maintain their shape (Fig. 3 and Fig. 4 A; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Recent results indicate that these opposing forces can lead to a transient and partial reduction of cell adhesion after mitotic exit. In Drosophila epithelia, E-cadherin levels are reduced at the interface between the cleavage furrow of the dividing cell and its neighbors (Fig. 3, B and C; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Specifically in embryonic epithelia, the local reduction of E-cadherin facilitates membrane separation, and a gap appears between the dividing cell and its neighbors (Fig. 3 B; Guillot and Lecuit, 2013a). In the dorsal thorax, in contrast, the neighbor cell plasma membrane detaches from the cortex and is drawn into the cleavage furrow (Fig. 3 C; Herszterg et al., 2013, 2014). What triggers E-cadherin modulation in cells after mitotic exit? The loss of overall cell polarity is one possible mechanism. During mitosis in Drosophila, follicular epithelial cells lose cortical enrichment of some apical polarity proteins (aPKC, Crumbs, and Bazooka/Par3; Bergstralh et al., 2013; Morais-de-Sá and Sunkel, 2013), and embryonic cells lose localization of lethal giant larvae, a basolateral cortical protein (Huang et al., 2009). Contrasting with these observations, however, MDCK cells and Drosophila embryonic and dorsal thorax epithelial cells appear to maintain apicobasal polarity as they divide (Reinsch and Karsenti, 1994; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Furthermore, E-cadherin reduction is limited to the furrow/neighbor interface and is not observed in other areas of cell contact. Therefore, an alternative mechanism that explains local E-cadherin modulation is mechanical tension that arises precisely at the area between the contractile ring and the neighboring cell membrane (Founounou et al., 2013; Guillot and Lecuit, 2013a).Does E-cadherin modulation serve a functional role in mitotic cells? In Drosophila embryonic and dorsal thorax epithelia, E-cadherin decrease leads to a local adhesion disengagement proposed to facilitate the formation of new AJs between daughter cells (Founounou et al., 2013; Guillot and Lecuit, 2013a). It has been previously reported that cells maintain their AJs throughout division. For example, intercellular junctions are maintained in dividing cells of human colonic mucosal crypt cells and basal keratinocytes (Baker and Garrod, 1993). Similarly, mitotic MDCK cultured cells maintain tight junctions apically and E-cadherin basolaterally (Reinsch and Karsenti, 1994). The E-cadherin loss in certain Drosophila epithelia may be either a tissue-specific phenomenon or a highly dynamic process only observable with the temporal resolution of live-cell imaging. Moreover, dividing cells in the Drosophila dorsal thorax show decreased levels of E-cadherin yet maintain their cohesiveness (Herszterg et al., 2013). Interestingly, E-cadherin is internalized in mitotic MDCK cells (Bauer et al., 1998). It will therefore be important to investigate whether loss of E-cadherin leads to adhesion disengagement in other epithelial tissues and whether tension alone or in combination with biochemical pathways is responsible for E-cadherin modulation.

Epithelial neighbors exert tension on daughter cell membranes to facilitate new AJ formation.

How new junctional contacts form during mitosis is a poorly understood problem at the heart of epithelial cell biology. In Drosophila, new membrane interfaces between nascent daughter cells initially show only a weak level of E-cadherin clusters (Guillot and Lecuit, 2013a; Herszterg et al., 2013). Subsequently, the daughter cells assemble their AJs de novo. How is the length of these new junctions determined with respect to cell geometry? Recent evidence indicates that AJ length is a function of local cell packing within the epithelium. In dividing cells of the Drosophila dorsal thorax, the contractile ring triggers tension and accumulation of myosin II in neighbors at the furrow/neighbor interface (Fig. 4 B; Founounou et al., 2013; Herszterg et al., 2013). Myosin II in the neighboring cells in turn contracts and creates tension at the furrowing membrane of the nascent daughter cells, keeping them tightly pressed against each other (Fig. 4, B and C). This local membrane juxtaposition facilitates AJ formation. To allow expansion of the daughter cell interface and maintain AJ length, branched actin polymerization via Rac1 and Arp2/3 is oriented to the midbody, which serves as a positional landmark for new AJs (Fig. 4 C; Herszterg et al., 2013). The midbody is a narrow intercellular bridge that remains after the contracted cytokinetic ring has driven membrane invagination, and it recruits the abscission factors that will eventually separate the daughter cells (Fededa and Gerlich, 2012). Interestingly, the midbody is positioned apically as a result of the presence of AJs. In Drosophila follicular epithelia, the midbody also provides cues for the formation of the apical daughter cell interface, suggesting that it plays a role in both AJ and epithelial cell polarity establishment and maintenance in dividing epithelial cells (Morais-de-Sá and Sunkel, 2013). Thus, examples from Drosophila epithelia show that cohesion between dividing cells and their neighbors together with the apically positioned midbody provides a spatial template and polarized positional cue for de novo AJ assembly (Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Further work on other epithelial tissues may provide alternative mechanisms of junction biogenesis.

Growth and order in the epithelium: Thinking outside the cell

During development, epithelial monolayers have the remarkable capacity to maintain specialized morphologies and barrier functions during rapid cell proliferation. Mitotic cells remain adherent to their neighbors throughout cell division. Cell cohesion enables local geometry and global tissue tension to instruct mitotic cells where to position their cleavage plane and how to assemble their junctions. However, local tension may also lead to a transient disengagement of dividing cells from their neighbors after mitotic exit. How is global and local tension conveyed to protein complexes in mitotic cells so that different outcomes take place? Moreover, it is unclear whether and how tissue tension instructs synchronously dividing epithelial cells how to divide and reestablish their junctions after division. Clearly, this is a fundamental problem for the maintenance of epithelial order and may be linked to the origin of epithelial cancers, in which cells undergo rapid proliferation but fail to remain integrated into the monolayer.The selected studies discussed here hint at the remarkable level of coordination that occurs during epithelial cell division, recasting mitosis as a truly multicellular process. Looking ahead, understanding the interface between cells, proteins, and mechanical forces that each operate on different scales will require creative multidisciplinary approaches in diverse organismal systems. Indeed, epithelial organization is widespread in nature and is encountered among even the most basal animals, including sponges and cnidarians as well as the fruiting body of the nonmetazoan social amoeba Dictyostelium discoideum (Wood, 1959; Ereskovsky et al., 2009; Houliston et al., 2010; Dickinson et al., 2011; Meyer et al., 2011). Combined, future interdisciplinary studies and a fresh look at diverse animal models should yield new insight into epithelial cell division for many years to come.     

Cell biology in neuroscience: Death of developing neurons: New insights and implications for connectivity

The concept that target tissues determine the survival of neurons has inspired much of the thinking on neuronal development in vertebrates, not least because it is supported by decades of research on nerve growth factor (NGF) in the peripheral nervous system (PNS). Recent discoveries now help to understand why only some developing neurons selectively depend on NGF. They also indicate that the survival of most neurons in the central nervous system (CNS) is not simply regulated by single growth factors like in the PNS. Additionally, components of the cell death machinery have begun to be recognized as regulators of selective axonal degeneration and synaptic function, thus playing a critical role in wiring up the nervous system.

Why do so many neurons die during development?

Programmed cell death occurs throughout life, as cell turnover is part of homeostasis and maintenance in most organs and tissues. The situation in the nervous system is principally different, as the vast majority of neurons undergo their last round of cell division early in development. Soon after exiting the cell cycle, neurons start elongating axons to innervate their targets. It is during this period that they are highly susceptible to undergo programmed cell death: a large percentage, as much as 50% in several ganglia in the peripheral nervous system (PNS) as well as in various central nervous system (CNS) areas, is eliminated around the time that connections are being made with other cells. Later in development, the propensity of neurons to initiate apoptosis progressively decreases. The likelihood for a neuron to undergo apoptosis seems to be determined by a tightly regulated apoptotic machinery (summarized in Fig. 1). Therefore, modulation of the expression levels or the activity of components of this apoptotic balance changes the sensitivity to death-promoting cues, allowing temporal restriction of cell death.Open in a separate windowFigure 1.Core components of the apoptotic machinery. The likelihood that a neuron undergoes apoptosis is determined by the interplay of the tightly interlinked apoptotic machinery, many components of which are highly conserved between species. The critical, and often terminal, step in programmed cell death is the proteolytic activation of the executor caspases (such as caspase 3, 6, 7) by the initiator caspases (i.e., caspase 8, 9, and 10; Riedl and Salvesen, 2007). In mammalian cells, initiation of the executor caspases is regulated by two distinct protein cascades: the intrinsic pathway, also known as the mitochondrial pathway, and the extrinsic pathway. The intrinsic pathway integrates a number of intra- and extracellular signal modalities, such as redox state (for example, the reactive oxygen species; Franklin, 2011), DNA damage (Sperka et al., 2012), ER stress (Puthalakath et al., 2007) and growth factor deprivation (Deckwerth et al., 1998; Putcha et al., 2003; Bredesen et al., 2005), or activation of the p75NTR neurotrophin receptor by pro-neurotrophins (Nykjaer et al., 2005). The stressors converge onto pro- and anti-apoptotic members of the Bcl-2 protein family (for example: BCL-2, BCL-X_l, BAX, and tBID; Youle and Strasser, 2008). These proteins regulate the release of cytochrome c from mitochondria, which activates the initiator caspase 9 through Apaf1 (Riedl and Salvesen, 2007). The extrinsic pathway links activation of ligand-bound death receptors (such as Fas/CD95 and TNFR) to the initiator caspase 8 and 10, through formation of the death-inducing signaling complex (DISC; LeBlanc and Ashkenazi, 2003; Peter and Krammer, 2003). Together with additional regulatory elements (including the Inhibitors of apoptosis proteins [IAP]; Vaux and Silke, 2005) and cFLIP (Scaffidi et al., 1999; Wang et al., 2005), the apoptotic machinery forms a balance that determines the propensity of the neuron to undergo apoptosis.Programmed cell death eliminates many neurons during development, even in organisms comprised of only few cells, such as Caenorhabditis elegans. As neurons and their targets are initially separated, it is possible that the initial generation of an overabundance of neurons is simply part of a mechanism to ensure that distal targets are adequately innervated (Oppenheim, 1991; Conradt, 2009; Chen et al., 2013). In various tissues other than the nervous system, programmed cell death is used to eliminate cells that are no longer needed, defective, or harmful to the function of the organism. However, there is strong evidence that the elimination of superfluous neurons in the developing nervous system is not essential. For example, early work in C. elegans revealed that preventing programmed cell death does not result in significant behavioral alterations (Ellis and Horvitz, 1986). In the C57BL/6 mouse strain, deletion of the executor caspases 3 and 7 (Fig. 1) has a remarkably limited neuropathological and morphological impact in the CNS (Leonard et al., 2002; Lakhani et al., 2006) compared with the 129X1/SvJ strain, in which deletion of these caspases causes neurodevelopmental defects (Leonard et al., 2002). Similar conclusions were reached by blocking the Bcl-2–associated X protein (BAX)–dependent pathway in many neuronal populations, including motoneurons (Buss et al., 2006a). A recent study in the developing retina showed that in mice lacking the central apoptotic regulator BAX, the normal mosaic distribution of intrinsically photosensitive retinal ganglion cells (ipRGCs) was perturbed (Chen et al., 2013). Although this abnormal distribution is dispensable for the intrinsic photosensitivity of the ipRGCs, it is required for establishing proper connections to other neurons in the retina, which is necessary for rod/cone photo-entrainment (Chen et al., 2013). Even though this finding highlights a physiological role for programmed cell death in the CNS, the functional consequences remain rather underwhelming in the face of a process that eliminates such large numbers of neurons (Purves and Lichtman, 1984; Oppenheim, 1991; Miller, 1995; Gohlke et al., 2004). It thus appears that apoptotic removal of the surplus neurons generated during development mainly serves the purpose to optimize the size of the nervous system to be minimal, but sufficient.

A molecular substrate for the neurotrophic theory

Quantitatively, programmed cell death of neurons in the PNS and CNS is most dramatic when neurons start contacting the cells they innervate. Because experimental manipulations such as target excision typically lead to the death of essentially all innervating neurons (Oppenheim, 1991), the concept emerged that the fate of developing neurons is regulated by their targets. This notion is often referred to as the “neurotrophic theory” (Hamburger et al., 1981; Purves and Lichtman, 1984; Oppenheim, 1991), but it is important to realize that it evolved in the absence of direct mechanistic or molecular support (Purves, 1988). Originally described as a diffusible agent promoting nerve growth, the eponymous NGF later provided a strong and very appealing molecular foundation for this theory (Korsching and Thoenen, 1983; Edwards et al., 1989; Hamburger, 1992). The tyrosine kinase receptor tropomyosin receptor kinase A (TrkA), which was initially identified as an oncogene (Martin-Zanca et al., 1986), was fortuitously discovered to be the critical receptor necessary for the prevention of neuronal cell death by NGF (Klein et al., 1991). Both the remarkable expression pattern of TrkA in NGF-dependent neurons and the onset of its expression during development (Martin-Zanca et al., 1990) provided further additional support for the neurotrophic theory. However, for a surprisingly long time, the question was not asked as to why only specific populations of neurons strictly depend on NGF for survival, while others do not. Indeed, it was only recently shown that TrkA causes cell death of neurons by virtue of its mere expression, and that this death-inducing activity is prevented by addition of NGF (Nikoletopoulou et al., 2010). These findings thus indicate that TrkA acts as a “dependence receptor,” a concept introduced after observations that various cell types die when receptors are expressed in the absence of their cognate ligands (Bredesen et al., 2005; Tauszig-Delamasure et al., 2007). Accordingly, embryonic mouse sympathetic or sensory neurons survive in the absence of NGF when TrkA is deleted (Nikoletopoulou et al., 2010). The closely related neurotrophin receptor TrkC also acts as a dependence receptor (Tauszig-Delamasure et al., 2007; Nikoletopoulou et al., 2010). Here, it is interesting to note a series of older, convergent results indicating that deletion of neurotrophin-3 (NT3), the TrkC ligand, leads to a significantly larger loss of sensory and sympathetic neurons in the PNS than the deletion of TrkC (Tessarollo et al., 1997). This phenotypic discrepancy fits well with the idea that inactivation of the ligand of a dependence receptor is expected to yield a more profound phenotype than inactivation of the receptor itself (Tauszig-Delamasure et al., 2007). How TrkA and TrkC induce apoptosis remains to be fully elucidated. It seems that proteolysis is involved, either of TrkC itself (Tauszig-Delamasure et al., 2007), as was suggested for other dependence receptors (Bredesen et al., 2005), or by the proteolysis of the neurotrophin receptor p75NTR, which associates with both TrkA and TrkC (Fig. 2; Nikoletopoulou et al., 2010). Surprisingly, although TrkA and TrkC cause cell death, the structurally related TrkB receptor does not (Nikoletopoulou et al., 2010), a difference that appears to be accounted for by their differential localization in the cell membrane. TrkA and TrkC colocalize with p75NTR in lipid rafts, whereas TrkB, which also associates with p75NTR (Bibel et al., 1999), is excluded from lipid rafts (Fig. 2; unpublished data). Interestingly, the transmembrane domains of TrkA and TrkC are closely related, and differ clearly from that of TrkB. It turns out that a chimeric protein of TrkB with the transmembrane domain of TrkA causes cell death, which can be prevented by the addition of the TrkB ligand brain-derived neurotrophic factor (BDNF; unpublished data). The suggestion that the lipid raft localization of TrkA and TrkC is important for their death-inducing function is in line with a number of reports indicating that certain apoptotic proteins preferentially localize in lipid rafts in the plasma membrane. After activation of the extrinsic apoptosis pathway, translocation of the activated receptors to lipid rafts in the membrane is required for assembling the death-inducing signaling complex (DISC; Davis et al., 2007; Song et al., 2007). Indeed, regulators of the extrinsic pathway (e.g., cFLIP; Fig. 1) prevent this translocation, explaining how they attenuate cell death induction (Song et al., 2007). Similarly, the localization of the dependence receptor DCC (deleted in colorectal cancer) in lipid rafts is a prerequisite for its pro-apoptotic activity in absence of its ligand, Netrin-1 (Furne et al., 2006).Open in a separate windowFigure 2.TrkA and TrkC as dependence receptors: mode of action and contrast with TrkB. All Trk receptors associate with the pan-neurotrophin receptor p75NTR (Bibel et al., 1999). A critical step in the induction of apoptosis by TrkA is the release of the intracellular death domain of p75NTR by the protease γ-secretase (Nikoletopoulou et al., 2010), which is localized in lipid rafts (Urano et al., 2005). Our membrane fractionation studies indicate that while TrkA and TrkC associate with p75NTR in lipid rafts, TrkB associated with p75NTR is excluded from this membrane domain (unpublished data). The 24–amino acid transmembrane domain of the Trk receptors may be responsible for this differential localization (see text).Despite the fact that TrkB does not act as a dependence receptor, its activation by BDNF is required for the survival of several populations of cranial sensory neurons (Ernfors et al., 1995; Liu et al., 1995). It appears that other death-inducing receptors predispose these neurons to be eliminated, such as p75NTR, which is expressed at high levels in some of these ganglia, or TrkC in vestibular neurons (Stenqvist et al., 2005). This latter case is of special interest, as NT3 is known not to be required for the survival of these neurons (Stenqvist et al., 2005). In addition to inducing apoptosis in the absence of their ligand, TrkA and TrkC have long been recognized to have a pro-survival function similar to TrkB, as can be inferred from the loss of specific populations of peripheral sensory neurons in mutants lacking these receptors (Klein et al., 1994; Smeyne et al., 1994).

Cell death in the CNS

Although TrkA is primarily expressed in peripheral sympathetic and sensory neurons, it is also found in a small population of cholinergic neurons in the basal forebrain (Sobreviela et al., 1994), a proportion of which requires NGF for survival (Hartikka and Hefti, 1988; Crowley et al., 1994; Müller et al., 2012). Selective deletion of TrkA was recently shown not to cause the death of these neurons (Sanchez-Ortiz et al., 2012). This supports the notion that TrkA acts as a dependence receptor for this small population of CNS neurons, like for peripheral sensory and sympathetic neurons. TrkA activation by NGF is essential for the maturation, projections, and function of these neurons (Sanchez-Ortiz et al., 2012), as was previously described for sensory neurons in the PNS as well (Patel et al., 2000).Whether or not receptors other than TrkA act as dependence receptors in the CNS is an important open question, particularly because TrkB, which is expressed highly by most CNS neurons, does not act as a dependence receptor (Nikoletopoulou et al., 2010). In retrospect, the structural similarities between TrkA and TrkB, just like those between NGF and BDNF (Barde, 1989), have substantially misled the field by suggesting that BDNF would act in the CNS like NGF in the PNS. Adding to the confusion were early findings showing that BDNF supports the growth of spinal cord motoneurons in vitro or in vivo after axotomy (Oppenheim et al., 1992; Sendtner et al., 1992; Yan et al., 1992). However, in the absence of lesion, deletion of BDNF does not lead to significant losses of neurons in the developing or adult CNS (Ernfors et al., 1994a; Jones et al., 1994; Rauskolb et al., 2010), unlike the case in some populations of PNS neurons. The poor correlation of the role of BDNF in CNS development and in axotomy and in vitro experiments is surprising, especially because the role of NGF in vivo could in essence be recapitulated by in vitro experiments. Although the reasons for this discrepancy are not fully understood, the strong up-regulation of death-inducing molecules such as p75NTR after axotomy (Ernfors et al., 1989) may be a part of the explanation. At present, most of the growth factors promoting the survival of PNS neurons fail to show significant survival properties for developing neurons in the CNS, as for example was shown for NT3 (Ernfors et al., 1994b; Fariñas et al., 1994), glial cell line–derived neurotrophic factor (GDNF; Henderson et al., 1994), ciliary neurotrophic factor (CNTF; DeChiara et al., 1995), and several others.In the developing CNS, neuronal activity and neurotransmitter input seem to play a more significant role than single growth factors in regulating neuronal survival. In particular, it has been known for a long time that blocking synaptic transmission at the neuromuscular junction has a pro-survival effect on spinal cord motoneurons (Pittman and Oppenheim, 1978; Oppenheim et al., 2008). By contrast, surgical denervation of afferent connections leads to increased apoptosis of postsynaptic neurons (Okado and Oppenheim, 1984), whereas inhibiting glycinergic and GABAergic synaptic transmission has both pro- and anti-apoptotic effects on motoneurons (Banks et al., 2005). Throughout the developing brain, blocking glutamate-mediated synaptic transmission involving NMDA receptors markedly increases normally occurring neuronal death (Ikonomidou et al., 1999; Heck et al., 2008). The mechanism involves a reduction of neuronal expression of anti-apoptotic proteins, such as B-cell lymphoma 2 (BCL-2; Hansen et al., 2004). Conversely, a limited increase in neuronal activity leads to down-regulation of the pro-apoptotic genes BAX and caspase 9 (Léveillé et al., 2010), thereby reducing the propensity of these cells to initiate programmed cell death (Hardingham et al., 2002). In addition to directly modulating the expression of apoptotic proteins, neuronal activity affects the expression of several secreted growth factors, such as BDNF (Hardingham et al., 2002; Hansen et al., 2004) and GDNF (Léveillé et al., 2010). So, even though BDNF is not a major survival factor in the developing CNS, it appears to be critical for activity-dependent neuroprotection (Tremblay et al., 1999). A recent publication revealed that certain populations of neurons in the CNS do not follow the predictions of the neurotrophic theory and showed that apoptosis of cortical inhibitory neurons is independent of cues present in the developing cerebral cortex (Southwell et al., 2012). This study indicates that programmed cell death of a large proportion of interneurons in the CNS is regulated by intrinsic mechanisms that are largely resistant to the presence or absence of extrinsic cues (Dekkers and Barde, 2013).Taken together, even though the extent of naturally occurring cell death in the different regions of the CNS is not nearly as well characterized as in the PNS, let alone quantified, it appears that its regulation may significantly differ. Although single secreted neurotrophic factors seem to be largely dispensable for survival, neuronal activity and other intrinsic mechanisms drive the propensity of the neurons in the CNS to undergo apoptosis. An important open question in this context is a possible involvement of non-neuronal cells, such as glial cells (see Corty and Freeman, in this issue).

The apoptotic machinery as a regulator of connectivity

Activation of the executor caspases has been most studied in cell bodies and typically results in the demise of the entire cell (Williams et al., 2006). However, recent evidence shows that caspases are also activated locally in neuronal processes and branches destined to be eliminated, for example in axons overshooting their targets that are subsequently pruned back to establish the precise adult connectivity (Finn et al., 2000; Raff et al., 2002; Luo and O’Leary, 2005; Buss et al., 2006b). Initially, axonal degeneration and axon pruning were thought to be independent of caspases (Finn et al., 2000; Raff et al., 2002). Later work in Drosophila melanogaster (Kuo et al., 2006; Williams et al., 2006) and in mammalian neurons (Plachta et al., 2007; Nikolaev et al., 2009; Vohra et al., 2010) demonstrated that interfering with the apoptotic balance or the executor caspases can prevent or at least delay axonal degeneration. Simon et al. (2012) have found that a caspase 9 to caspase 3 cascade is crucial for axonal degeneration induced by NGF withdrawal, with caspase 6 activation playing a significant but subsidiary role. Upstream of the caspases, BCL-2 family members such as BAX and BCL-X_l are required (Nikolaev et al., 2009; Vohra et al., 2010). It is conceivable that the failure of ipRGCs in BAX-deficient mice to form appropriate connections to other cells in the retina (Chen et al., 2013) may be in part attributable to defective axonal degeneration. Surprisingly, Apaf1 appears not to be involved in this process (Cusack et al., 2013), suggesting that axon degeneration depends on the concerted activation of the intrinsic initiator complex in a different way from apoptosis.Strikingly, a series of recent studies showed that several caspases and components of the intrinsic pathway also affect normal synaptic physiology in adulthood (Fig. 3, A–D). Here, pro-apoptotic proteins are predominantly involved in weakening the synapses, whereas the anti-apoptotic proteins have been mainly associated with synaptic strengthening (Fig. 3 B). In particular, caspase 3 promotes long-term depression (LTD), a stimulation paradigm that results in a period of decreased synaptic transmission (Li et al., 2010), and also prevents long-term potentiation (LTP), the converse situation leading to strengthened synaptic transmission (Jo et al., 2011). Likewise, the proapoptotic BCL-2 family members BAX and BAD stimulate LTD (Jiao and Li, 2011). By contrast, the anti-apoptotic protein BCL-X_l increases synapse numbers and strength (H. Li et al., 2008), and the inhibitor of apoptosis protein (IAP) family member survivin was reported to be involved in LTP in the hippocampus (Iscru et al., 2013) and in activity-dependent gene regulation (O’Riordan et al., 2008).Open in a separate windowFigure 3.Canonical and noncanonical functions of the apoptotic machinery. (A) The apoptotic machinery is not only involved in eliminating cells destined to die, but is also a central player in refining neuronal connectivity, by regulating synaptic transmission and by generating the adult connectivity through axon pruning (Luo and O’Leary, 2005; Hyman and Yuan, 2012). But how the canonical and noncanonical roles of the apoptotic machinery are interlinked and spatially restricted is not well understood. (B) In the adult nervous system, the pro-apoptotic proteins BAX, caspase 9, and caspase 3 promote weakening of synapses (long-term depression [LTD]; Li et al., 2010; Jiao and Li, 2011; Jo et al., 2011), while the anti-apoptotic proteins Bcl-X_l and the IAP survivin promote synaptic strengthening (long-term potentiation [LTP]; Li et al., 2008a; Iscru et al., 2013). It is unclear how the activation of these pathways is restricted to a single synapse, but a recent review suggested that the proteasomal degradation of activated caspases may prevent their diffusion (Hyman and Yuan, 2012). (C) Caspase activation is now known to be required for axon pruning during development to generate the adult refined connectivity (Luo and O’Leary, 2005; Simon et al., 2012). Different pathways are activated depending on the stimulus leading to degeneration. Growth factor deprivation during development leads to activation the executor caspases 3 and 6 (Simon et al., 2012) through the intrinsic apoptotic pathway, although its core protein Apaf1 does not seem to be required for this process (Cusack et al., 2013). On the other hand, a traumatic injury leads to reduced influx of NMNAT2 into the axon, which negatively affects the stability and function of mitochondria and leads to an increased calcium concentration (Wang et al., 2012). The effector caspase, caspase 6, is dispensable for this form of axonal degeneration (Vohra et al., 2010; Simon et al., 2012). Regulatory proteins such as the IAPs and also the proteasome seem to play a role in limiting the extent of activation to the degenerating part of the axon (Wang et al., 2012; Cusack et al., 2013; Unsain et al., 2013). (D) Simplified schematic of the main pro- and anti-apoptotic components. DISC, death-induced signaling complex. IAP, inhibitor of apoptosis protein. See Fig. 1 for details.These findings indicate that the apoptotic machinery acts at different levels in the cell, ranging from driving sub-lethal degradation of a compartment (Fig. 3 C) and attenuating synaptic transmission at the neuronal network level (Fig. 3 B) to destroying the entire cell during development or in disease (Fig. 3 D). How the cell spatially restricts the extent of activation of the apoptotic machinery is yet unclear. For example, elimination of the somata of developing neurons after neurotrophin deprivation is preceded by axonal degeneration, but not all instances of axonal degeneration lead to the death of the neuron (Campenot, 1977; Raff et al., 2002). Local regulation of caspase activation by IAPs is well established as a means for ensuring the elimination of neuronal processes in D. melanogaster (Kuo et al., 2006; Williams et al., 2006). Recent findings suggest a similar role for IAP in mammalian neurons, where it limits caspase activation to the degenerating axon (Fig. 3 C; Cusack et al., 2013; Unsain et al., 2013). The spontaneous mutation Wallerian degeneration slow (WldS; Lunn et al., 1989) has been instrumental to understand that trauma-induced axon degeneration is a regulated process different from, and independent of, cell body degeneration (Wang et al., 2012), but also distinct from axon pruning (Hoopfer et al., 2006). Work on the chimeric protein encoded by the WldS mutation also led to the identification of the protein NMNAT2 (nicotinamide mononucleotide adenylyltransferase 2) as a labile axon survival factor (Gilley and Coleman, 2010). How the WldS chimeric protein and NMNAT2 result in axon protection is unclear, but several lines of evidence seem to converge on local regulation of mitochondrial function and motility (Avery et al., 2012; Fang et al., 2012).Related to the spatial limiting of apoptotic activity is the question of how a local source of neurotrophins leads to the rescue of a developing peripheral neuron. When neurons encounter a source of neurotrophins, only the receptors close to the target will be activated, whereas the others, located further away, are not. The cell, therefore, needs to integrate a pro-survival signal from the activated receptors, and death-inducing signals from the nonactivated dependence receptors. The continued signaling of activated neurotrophin receptors that are retrogradely transported to the soma (Grimes et al., 1996; Howe et al., 2001; Wu et al., 2001; Harrington et al., 2011) likely play a role in counteracting the pro-apoptotic signaling proximal to the source of neurotrophins. It will be interesting to investigate whether similar mechanisms play a role in axon pruning and traumatic axon degeneration as well.

Programmed cell death in the adult brain

Most of the nervous system becomes post-mitotic early in development. In rodents, two brain areas retain the capacity to generate new neurons in the adult: the sub-ventricular zone, which generates neurons that migrate toward the olfactory bulb, and the sub-granular zone of the dentate gyrus of the hippocampus, where neurons are generated that integrate locally. Similar to what is observed during embryonic development, these adult-generated neurons are produced in excess, and a large fraction undergoes apoptosis when contacting its designated targets (Petreanu and Alvarez-Buylla, 2002; Kempermann et al., 2003; Ninkovic et al., 2007). Preventing apoptosis of adult-generated neurons in the olfactory bulb only has limited functional consequences (Kim et al., 2007), whereas a similar maneuver in the dentate gyrus does lead to impaired performance in memory tasks (Kim et al., 2009). Why superfluous hippocampal neurons would need to be eliminated for proper function is a matter of speculation, but may be linked with the fact that these are excitatory projection neurons, whereas in the olfactory bulb only axon-less inhibitory granule cells are integrated. The extent of survival in both these areas critically depends on the activity of the neuronal network in which these newly born neurons have to integrate (Petreanu and Alvarez-Buylla, 2002; Kempermann et al., 2006; Ninkovic et al., 2007). In this context, BDNF, the expression level of which is well known to be regulated by network activity, supports the survival of young adult–generated neurons and possibly even stimulates the proliferation of neural progenitors (Y. Li et al., 2008; Waterhouse et al., 2012). Interestingly, in young adult mouse mutants that exhibit spontaneous epileptic seizures, significantly higher levels of BDNF have been measured (Lavebratt et al., 2006; Heyden et al., 2011). Concomitantly, the entire hippocampal formation is considerably enlarged by as much as 40% (Lavebratt et al., 2006; Angenstein et al., 2007), which in turn is dependent on the epileptic seizures (Lavebratt et al., 2006). Whether or not there is a causal relationship between increased BDNF levels and hippocampal volume remains to be established.

Conclusion

Now that is has become clear that action of the apoptotic machinery can be limited spatially and temporally, several questions need to be addressed: how do neurons integrate intrinsic and extrinsic pro- and anti-apoptotic signals; and how they are spatially restricted to allow degradation of a dendrite or axon, or modulation of synaptic transmission? Another important issue is the regulation of cell death by intrinsic mechanisms in the central nervous system of vertebrates, not least because programmed cell death is observed in the CNS in a number of neurodegenerative diseases (Vila and Przedborski, 2003). Indeed, several of the central apoptotic components discussed here are also involved in these disorders (Hyman and Yuan, 2012). New insights in the regulation of programmed cell death in the developing nervous system may therefore continue to help to better understand the pathophysiological mechanisms of neurodegenerative disorders.     

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.     

Aneuploidy in health,disease, and aging

Synapse formation is a highly regulated process that requires the coordination of many cell biological events. Decades of research have identified a long list of molecular components involved in assembling a functioning synapse. Yet how the various steps, from transporting synaptic components to adhering synaptic partners and assembling the synaptic structure, are regulated and precisely executed during development and maintenance is still unclear. With the improvement of imaging and molecular tools, recent work in vertebrate and invertebrate systems has provided important insight into various aspects of presynaptic development, maintenance, and trans-synaptic signals, thereby increasing our understanding of how extrinsic organizers and intracellular mechanisms contribute to presynapse formation.Chemical synapses are highly specialized, asymmetric intercellular junction structures that are the basic units of neuronal communication. Proper development of synapses determines appropriate connectivity for the assembly of functional neuronal circuits. Synaptic circuits arise during development through a series of intricate steps (Waites et al., 2005; McAllister, 2007; Jin and Garner, 2008). First, spatiotemporal cues guide axons through complex cellular environments to contact their appropriate postsynaptic targets. At their destination, synapse formation is specified and initiated through adhesive interactions between synaptic partner cells or by local diffusible signaling molecules. Stabilization of intercellular contacts and assembly into functional synapses involves cytoskeletal rearrangements, aggregation, and insertion of pre- and postsynaptic components at nascent synaptic sites. Maturation and modulation of these newly formed synapses can then occur by altering the organization or composition of synaptic proteins and post-translational modifications to achieve its required physiological responsiveness (Budnik, 1996; Lee and Sheng, 2000). Conversely, retraction of contacts and elimination of inappropriate synaptic proteins help to refine the neuronal circuitry (Goda and Davis, 2003; Sanes and Yamagata, 2009).Over the last decade, new insights have furthered our understanding of synapse development through the identification of new molecular players and by advanced imaging technology that has allowed for high-resolution inspection of the dynamics and relative positions of synaptic proteins. This review will highlight recent results on the development of presynaptic specializations, and the roles of trans-synaptic organizers, intracellular synaptic proteins, and the cytoskeleton during the formation and maintenance of synapses.

Axonal transport of synaptic vesicle and active zone proteins

After cell fate determination and morphogenesis, neurons continue to differentiate by entering the phase of synapse formation. Most synaptic material required for this process is synthesized in the cell body of neurons and transported to synapses by microtubule (MT)-based molecular motors (Fig. 1). MTs are intrinsically polarized filaments with a plus and a minus end (Fig. 1 B). MT-based molecular motors use this polarity to transport cargoes to specific cellular locations. Examination of MTs by electron microscopy in dissociated cultured neurons showed that the organizations of MTs is different in axon and dendrite (Baas et al., 1988, 2006). In axons, all microtubules have their minus ends oriented toward the cell body and their plus ends extend distally. On the contrary, the MT polarity in dendrites is mixed. Recent studies tracking the movement of end-binding MT-capping proteins confirmed these results in vivo. Specifically, axonal MTs are uniformly organized with their plus ends pointing distally in all organisms. Dendrites of vertebrate neurons show more plus end–out MTs in vivo, whereas flies and worms have more minus end–out MTs in dendrites (Stepanova et al., 2003; Rolls et al., 2007; Stone et al., 2008).Open in a separate windowFigure 1.Regulatory steps during polarized motor-based transport of synaptic material. (A) At the Golgi apparatus, synaptic proteins have to be sorted into appropriate vesicles. These vesicles and other cargo such as mitochondria get loaded onto specific motor proteins. (B) Establishment of proper microtubule polarity along the axon determines anterograde and retrograde trafficking by plus end– and minus end–directed motor proteins such as kinesins and dynein. (C) At the appropriate destination, motor-cargo unloading occurs in a regulated fashion to achieve the appropriate distribution of synaptic boutons. At synapses, synaptic vesicle precursors give rise to mature synaptic vesicles. Proteins required for the SV cycle and trans-synaptic adhesion coalesce into the active zone (AZ) underneath the plasma membrane juxtaposed against the postsynaptic membrane.Does the difference in microtubule organization and polarity help to segregate synaptic cargoes between axons and dendrites? Recent studies have started to identify some molecules that create these differences in MT polarity in different neuronal subcellular compartments and show how disruption of their function affects synapse formation. For example, a recent paper showed that kinesin-1 is required to establish the predominantly minus end–out organization in the dendrites of Caenorhabditis elegans motor neurons (Yan et al., 2013). In kinesin-1/unc-116 mutants, dendrites adopt the axon-like MT polarity causing presynaptic cargoes to mislocalize into dendrites (Seeger and Rice, 2010; Yan et al., 2013). Similarly, loss of the MT-binding CRMP protein UNC-33 or the actin–spectrin adaptor protein ankyrin/UNC-44 in worms also results in MT polarity defects, which also results in ectopic localization of synaptic vesicles and active zone proteins into dendrites (Maniar et al., 2012). These results support the idea that MT polarity ensures the faithful targeting of presynaptic components to the axon. However, another way motors can distinguish between axons and dendrites is through MT-associated proteins (MAPs). In a recent study, Banker and colleagues showed that plus end–orienting kinesins can differentiate axon and dendrite, likely due to specific MT-binding proteins in these compartments (Huang and Banker, 2012).The direct regulation of motor activity by MTs or synaptic vesicle–associated proteins is likely to contribute to the trafficking of synaptic cargoes. Doublecortin, a MAP, binds to kinesin-3/KIF1A to affect the trafficking of the synaptic vesicle protein, synaptobrevin, in hippocampal neurons by altering the affinity of ADP-bound KIF1A to MTs (Liu et al., 2012). The Rab3 guanine nucleotide exchange factor, DENN/MADD, functions as an adaptor between kinesin-3 and GTP-Rab3–containing synaptic vesicles to promote the trafficking of synaptic vesicles in the axon (Niwa et al., 2008).Precise regulation of motor-based transport ensures that synaptic cargoes are delivered to and maintained at synapses. Several recent studies have provided evidence that two postmitotic cyclin-dependent kinases are important regulators of anterograde and retrograde trafficking of presynaptic cargoes. The kinase CDK-5 is required in many aspects of nervous system function. In the context of presynaptic development and function, CDK-5 has been shown to regulate the transport of synaptic vesicles and dense core vesicles, which contain neuropeptides, by inhibiting a dynein-mediated pathway that mobilizes presynaptic components to the somatodendritic compartments in C. elegans neurons (Ou et al., 2010; Goodwin et al., 2012). A paralogue of CDK-5, the PCT-1 kinase acts in a partially redundant pathway to prevent the mislocalization of presynaptic material to dendrites. In animals lacking both kinases or their activators, synaptic cargoes completely mislocalize to the dendrites, leaving an “empty” axon (Ou et al., 2010). Vertebrate CDK-5 also plays profound roles in the regulation of synaptic vesicle pools by modifying Ca²⁺ channels. Genetic ablation or pharmacological inhibition of CDK-5 increases the pool of synaptic vesicles that are docked at the active zone, termed the readily releasable pool, and potentiates synaptic function (Kim and Ryan, 2010, 2013). These results suggest that CDK-5 and its paralogue control local and global vesicle pools. Regulation of the exchange between these pools can affect membrane trafficking at presynaptic terminals as well as the overall polarity of neurons.To form synapses at defined locations, cargoes not only need to know how to “get on” the transport system but also need to know where to precisely “get off” at their destination (Fig. 1 C). Loss of a conserved small G-protein of the Arf-like family, ARL-8, in C. elegans, resulted in premature exit of synaptic cargoes during transport and showed ectopic aggregations of synaptic vesicles in the proximal axon. This causes a reduction in the number but an increase in the size of synapses (Klassen et al., 2010). ARL-8 localizes to both stable and trafficking synaptic vesicles and promotes trafficking by increasing kinesin-3 activity and suppressing aggregation-induced stoppage of synaptic cargoes along the axon (Wu et al., 2013). Hence, the balance between motor activity and aggregation propensity of trafficking cargoes may determine the number, size, and location of presynaptic terminals. Interestingly, the small GTPase Rab3, which normally associates with synaptic vesicles, has recently been shown to affect the distribution of active zone proteins at fly neuromuscular junction (NMJ) synapses, further suggesting that the trafficking of synaptic vesicles and formation of active zones are linked (Graf et al., 2009).Besides synaptic material, another major organelle cargo that is often present at the presynaptic terminal is mitochondria. The Milton–Miro complex functions as an adaptor between kinesin-1 and mitochondria to support axonal transport of mitochondria. Interestingly, the coupling of the Milton–Miro complex to kinesin is regulated by Ca²⁺ (Macaskill et al., 2009; Wang and Schwarz, 2009), providing a mechanism for neuronal activity controlling transport of mitochondria along the axon.Previous studies have suggested that components of the presynaptic active zone are transported in a preassembled form by Piccolo-Bassoon transport vesicles (PTVs) that may contain multiple components required to build a synapse (Zhai et al., 2001; Shapira et al., 2003). Recent studies found that Golgi-derived PTVs contain many active zone proteins including Piccolo, Bassoon, RIM1α, and ELKS2/CAST, but lack another active zone component, Munc-13, which may exit the Golgi on separate vesicles (Maas et al., 2012). Packing of various active zone components that have the propensity to self-assemble into separate vesicles may contribute a way to control synaptogenesis. This is interesting in light of the finding that Munc-13 can function as a protein scaffold for Bassoon and ELKS2 (Wang et al., 2009). The link between trafficking of synaptic vesicle and active zone components is not well understood. In vivo time-lapse imaging of synaptic vesicle and active zone trafficking showed that these components, possibly in the form of dense core vesicles, could be trafficked together in C. elegans neurons, suggestive of prepackaged presynaptic material during transport (Wu et al., 2013). Taken together, axonal transport of synaptic components is a necessary step for synapse formation and maintenance. The regulation of MTs, molecular motors, and synaptic cargoes ensure the targeting of appropriate proteins to synapses.

Role of the actin cytoskeleton in presynaptic assembly

Although MT-mediated transport is critical for long-range trafficking, actin-based mechanisms often organize local protein complexes in subcellular domains. A large body of work has described the role of the actin cytoskeleton in postsynaptic structure and function (Schubert and Dotti, 2007; Hotulainen and Hoogenraad, 2010). We will focus on more recent work that has highlighted the importance of the actin cytoskeleton in presynaptic formation.F-actin is required for presynaptic assembly during the early stages of synaptogenesis. Depolymerization of F-actin in young hippocampal neuronal cultures results in a reduction in the size and number of synapses. This effect was not seen with older cultures when synapses are more mature (Zhang and Benson, 2001). This observation correlates with an increase in both pre- and postsynaptic F-actin levels in newly formed synapses compared with mature synapses (Zhang and Benson, 2002).F-actin has been implicated in many steps of synapse assembly and function (Fig. 2; Cingolani and Goda, 2008). One of the roles that has been proposed for F-actin is to act as a scaffold for other presynaptic proteins (Sankaranarayanan et al., 2003). A recent study identified an F-actin–binding active zone molecule Neurabin/NAB-1 that is recruited by a presynaptic F-actin network (Chia et al., 2012). In addition, knockdown of Rac/Cdc42 GTPase exchange factor β-Pix resulted in a decrease in actin at synapses with a concomitant loss of synaptic vesicle clustering (Sun and Bamji, 2011). These studies demonstrate that F-actin at presynaptic sites can recruit and stabilize presynaptic components.Open in a separate windowFigure 2.Assembling the presynaptic active zone. Scaffolding proteins including Liprin, SYD-1, ELKS, Neurabin, Piccolo, and Bassoon form the dense protein network in the presynaptic cytomatrix that facilitates synaptic vesicle docking and fusion. The presynaptic F-actin networks are required for presynaptic assembly and maintenance.Studies of Drosophila NMJs have found that the presynaptic spectrin–actin cytoskeleton is important for synapse stability. Loss of presynaptic spectrin led to retraction of synapses (Pielage et al., 2005). Intriguingly, loss of postsynaptic spectrin increased the total number of the active zone specializations, termed T-bars, and affected the size and distribution of presynaptic sites. Thus, the spectrin cytoskeleton can impose a trans-synaptic influence on synapse development (Pielage et al., 2006).Given the importance of F-actin at synapses, it is crucial to understand the signaling pathways that instruct F-actin organization. Multiple studies have shown that signaling from synaptic cell adhesion molecules can lead to cytoskeletal rearrangements at synapses. Adhesion of hippocampal neurons to syndecan-2–coated beads is sufficient to induce F-actin clustering and downstream formation of presynaptic boutons (Lucido et al., 2009). In mice, the adhesion molecule L1CAM may bind to spectrin–actin adaptor ankyrin to mediate GABAergic synapse formation (Guan and Maness, 2010). Another adhesion molecule of the immunoglobulin superfamily SYG-1 in C. elegans has also been shown to be necessary and sufficient to recruit F-actin to synapses (Chia et al., 2012). In a recent study, secreted bone morphogenetic protein (BMP) can signal in a retrograde fashion to regulate Rac-GEF Trio expression in presynaptic neurons, which is important for controlling synaptic growth (Ball et al., 2010).Interestingly, presynaptic active zone proteins can also affect F-actin assembly (Fig. 2). Knockdown of Piccolo reduced activity-dependent assembly of F-actin at synapses and enhanced dispersion of Synapsin1a and synaptic vesicles in hippocampal neurons. Loss of Piccolo also resulted in a loss of Profilin 2, a regulator of actin polymerization (Waites et al., 2011).Various studies have begun to shed light on the actin regulators required for synaptic F-actin establishment and maintenance. Diaphanous, a formin-related gene that associates with barbed ends of F-actin, was found to function downstream of presynaptic receptor Dlar at fly NMJs. Spectrin–actin capping protein, Adducin, is enriched at presynaptic sites and is required to prevent synapse retraction and elimination (Bednarek and Caroni, 2011; Pielage et al., 2011). Activators of the Arp2/3 complex, WASP and WAVE, have also been implicated in the regulation of F-actin at synapses (Coyle et al., 2004; Stavoe et al., 2012; Zhao et al., 2013). This diversity of F-actin modulators suggests that there are probably different F-actin structures at different stages of development or even in subcellular domains within the synapse. This is supported by observations that F-actin can localize with synaptic vesicles, at the active zone and in the perisynaptic region (Bloom et al., 2003; Sankaranarayanan et al., 2003; Waites et al., 2011; Chia et al., 2012). Thus, much remains to be done in our understanding how distinct F-actin structures are formed and regulated to mediate various processes during synapse assembly and maintenance.

Assembly of the molecular network at presynaptic terminals

Although F-actin might help to initiate the presynaptic assembly process, many other ensuing molecular interactions are required to form the mature presynaptic apparatus (Fig. 2). The presynaptic active zone is comprised of a framework of scaffolding proteins that function as protein-binding hubs for other presynaptic components. Piccolo and Bassoon are important vertebrate multidomain proteins that traditionally have been widely used as active zone markers. Recent electrophysiology data on Piccolo mutant and Bassoon knockdown neurons showed that these molecules are dispensable for synaptic transmission but affect synaptic vesicle clustering (Mukherjee et al., 2010). Furthermore, Piccolo and Bassoon were found to be required for maintaining synapse integrity by regulating ubiquitination and degradation of presynaptic components (Waites et al., 2013).Forward genetic approaches in worms and flies have made important contributions to our understanding of the presynaptic cytomatrix. Studies have found that two active zone scaffolding molecules, SYD-1 and Liprin-α/SYD-2, are required for proper synapse formation (Zhen and Jin, 1999; Patel et al., 2006; Astigarraga et al., 2010; Owald et al., 2010; Stigloher et al., 2011). Interestingly, at fly NMJs, SYD-1 is necessary for clustering presynaptic neurexin that in turn clusters postsynaptic neuroligin (Owald et al., 2012). The presynaptic assembly function of SYD-1 and SYD-2 appears to be conserved because mutation analysis of mammalian SYD-1 and knockdown of Liprin-α both caused defects in presynaptic development and function (Spangler et al., 2013; Wentzel et al., 2013). In flies, the active zone T-bar structure is comprised of ERC/CAST family protein bruchpilot (brp) as the major active zone organizing protein (Fouquet et al., 2009). Brp is not only present at the active zone but also plays important scaffolding roles in localizing Ca²⁺ channels. In C. elegans, the Brp homologue ELKS-1 is also localized to the active zone; however, the importance of ELKS-1 during development of synapses was only revealed in sensitized genetic backgrounds (Dai et al., 2006; Patel and Shen, 2009), suggesting that there are likely redundant molecular pathways for presynaptic assembly. In the vertebrate system, loss of one of the three ELKS genes, surprisingly, caused an increase in the inhibitory synaptic transmission (Kaeser et al., 2009). Besides Brp, Rab3-interacting molecule (RIM) binding protein (RBP) was found to be important for active zone structural integrity in flies. Using super-resolution microscopy, RBP was found to surround Ca²⁺ channels at T-bars and loss of RBP resulted in defective Ca²⁺ channel clustering and reduced evoked neurotransmitter release (Liu et al., 2011).Assembly of the presynaptic active zone is subjected to several layers of regulation. The assembly process is balanced by inhibitory mechanisms that control the number and size of synapses. Loss of the E3 ubiquitin ligase Highwire/RPM-1 results in an increased number of synaptic boutons in flies and multiple active zones in worms (Wan et al., 2000; Zhen et al., 2000). Working together with F-box protein FSN-1, RPM-1 down-regulates the DLK MAP kinase signaling pathway (Liao et al., 2004; Nakata et al., 2005; Yan et al., 2009). Another E3 ubiquitin ligase, the SKP complex, has been shown to eliminate transient synapses during development in worms (Ding et al., 2007). Therefore, ubiquitin-mediated mechanisms play important roles in controlling the presynaptic assembly program.Other inhibitory mechanisms include SRPK79D, a serine–arginine protein kinase discovered in flies that represses T-bar formation (Johnson et al., 2009). In the mutant, the T-bar component Brp is ectopically accumulated in the axonal shaft. Regulator of synaptogenesis, RSY-1, limits the extent of presynaptic assembly by directly binding to active zone scaffold molecule Liprin-α/SYD-2 and SYD-1 (Patel and Shen, 2009). In addition, Liprin-α/SYD-2 may inhibit its own activity via intramolecular interactions (Taru and Jin, 2011; Chia et al., 2013).Taken together, the presynaptic assembly process driven by scaffolding molecules is controlled by complex inhibitory mechanisms to achieve the appropriate extent of aggregation in the process of synapse formation.

Trans-synaptic signals orchestrate pre- and postsynaptic formation

Coordinated pre- and postsynaptic development requires the precise apposition of presynaptic components to postsynaptic specializations. It is conceivable that signals from pre and postsynaptic sides functioning across the synaptic cleft coordinate synaptic differentiation reciprocally. Although a vast assortment of factors have been identified as synaptic organizers, the fact that genetic ablation of some synaptic organizers in vivo fails to elicit dramatic synaptic defects suggests the incomplete view of the trans-synaptic signaling. Moreover, the underlying mechanisms and the cross talk of these signaling pathways are still unclear. In recent years, an emerging body of literature has begun to shed light on trans-synaptic signaling and the importance of environmental cues in synapse formation.

Adhesion proteins instruct synaptic differentiation

A large body of literature suggests that trans-synaptic interactions between synaptic adhesion molecules function bi-directionally for synapse formation and maturation (Fig. 3). Neurexin–neuroligin is the first pair to be shown to induce pre- and postsynapse formation (Scheiffele et al., 2000; Graf et al., 2004; Chih et al., 2005; Nam and Chen, 2005; Chubykin et al., 2007). Recent in vitro studies have unveiled more components interacting with neurexin or neuroligin in specific synaptic differentiation events (Fig. 3, B and C). In early developmental stages, a secreted synaptic organizer, thrombospondin 1 (TSP1, see next section) increases the speed of synaptogenesis through neuroligin 1 (Xu et al., 2010). At excitatory synapses, a retrograde signaling controls synaptic vesicle clustering, neurotransmitter release, and presynaptic maturation by cooperation of neuroligin and N-cadherin (Wittenmayer et al., 2009; Stan et al., 2010; Aiga et al., 2011). A leucine-rich repeat transmembrane (LRRTM) protein family was also identified as an organizer of the function of excitatory synapses through interactions with neurexin (Linhoff et al., 2009). Further studies showed that binding of LRRTMs and neuroligins to neurexin acts redundantly to maintain excitatory synapses by preventing activity and Ca²⁺-dependent synapse elimination during early development, while performing divergent functions upon synapse maturation (de Wit et al., 2009; Ko et al., 2009, 2011; Soler-Llavina et al., 2011).Open in a separate windowFigure 3.Adhesive trans-synaptic signalings orchestrate excitatory and inhibitory synaptic assembly. Multiple pairs of trans-synaptic adhesion molecules organize synaptic differentiation and function on both pre- and postsynaptic sites. Note that different adhesion molecules are used at excitatory and inhibitory synapses. LPH1, latrophilin 1; α-DG, α-dystroglycan; β-DG, β-dystroglycan; S-SCAM, synaptic scaffolding molecule; Lasso, LPH1-associated synaptic surface organizer; IL-1RAcp, interleukin-1 receptor accessory protein.The function of neurexin and neuroligin in mediating synaptic differentiation has also been shown at Drosophila NMJs and mammalian CNS. In mammalian, although neither compound knockout of three neurexins nor two individual neuroligin knockout mice display severe defects in the number or morphology of synapses (Missler et al., 2003), the deletion of either neurexin or neuroligin affects the neurotransmitter release and in turn impairs the relevant behavior (Zhang et al., 2005; Blundell et al., 2009, 2010; Etherton et al., 2009; Jedlicka et al., 2011). Neurexin loss of function in fly leads to reduced number and defective morphology of synaptic boutons and active zones from early developmental stages (Li et al., 2007; Chen et al., 2010). In contrast, deletion of either neuroligin 1 or 2 causes NMJ defects and alternations of active zones only in the larval stage, indicating that they function mainly in the expansion of NMJs during development (Banovic et al., 2010; Sun et al., 2011). These abnormalities further impair synaptic transmission at the NMJs (Li et al., 2007; Banovic et al., 2010; Chen et al., 2010; Sun et al., 2011). Moreover, these phenotypes are enhanced when the Teneurin family of adhesion molecules is deleted, suggestive of functional redundancy between adhesion molecules (Mosca et al., 2012). Recently, it has been reported that an active zone protein, SYD-1, is required for the formation and function of the neurexin–neuroligin complex in flies (Fig. 2; Owald et al., 2012), providing an example of how trans-synaptic neurexin–neuroligin signaling orchestrates synaptic assembly bi-directionally. Interestingly, at postsynaptic sites, the NMDA receptor activity-triggered Ca²⁺-dependent cleavage of neuroligin 1 was found to destabilize presynaptic neurexin, reduce presynaptic release probability, and depress synaptic transmission (Peixoto et al., 2012). This observation raises a possibility that neurexin and neuroligin could fine-tune synaptogenesis both positively and negatively.Although Drosophila neuroligin and neurexin mutants share many phenotypes in synaptic differentiation, there are some unique features for each mutant, suggesting that they play distinct roles. For example, some aspects of synaptic specificity are achieved by different pairs of neurexin–neuroligin interactions. Neuroligin 1 promotes the growth and differentiation of excitatory synapses by binding to PSD-95, whose amount balances the ratio of excitatory-to-inhibitory synaptic specializations (Prange et al., 2004; Banovic et al., 2010). Neuroligin 2, on the contrary, binds to a scaffold protein gephyrin at inhibitory synapses, instructing inhibitory postsynaptic assembly (Fig. 3, B and C; Poulopoulos et al., 2009). Different isoforms of neurexin also contribute to the differentiation of excitatory and inhibitory synapses (Fig. 3, B and C; Chih et al., 2006; Graf et al., 2006; Kang et al., 2008).Other novel trans-synaptic interactions have also been identified to organize synaptic differentiation (Fig. 3, B and C). For example, Netrin-G ligand 3 (NGL-3), localized at postsynaptic region, induces excitatory synaptic differentiation by interacting with the receptor tyrosine phosphatase LAR family proteins, including PTPδ and PTPσ (Woo et al., 2009; Kwon et al., 2010). PTPδ can also trans-interact with Slitrk3 and IL-1 receptor accessory protein (IL-1RAcP) to promote presynaptic formation (Takahashi et al., 2012; Yoshida et al., 2012). Molecules that function in other neuronal developmental processes have also been shown to regulate synaptic differentiation. Farp1, essential for the dynamics of dendritic filopodia, regulates postsynaptic development and triggers a retrograde signal promoting active zone assembly by binding to SynCAM 1 (Cheadle and Biederer, 2012). Teneurins, instructing synaptic partner selection in fly olfactory system (Hong et al., 2012), act in synaptogenesis through trans-synaptic interaction at NMJs (Mosca et al., 2012). Another splice variant of a postsynaptic Teneurin-2 in rat, Lasso, binding with presynaptic Latrophilin 1 (LPH1), induces presynaptic Ca²⁺ signals and regulates synaptic function (Silva et al., 2011). Neural activity is also involved in controlling the growth of the presynapse. Conditioning or BDNF application induces presynaptic bouton development via an ephrin-B–dependent manner (Li et al., 2011), suggesting the role of EphB/ephrin-B signaling in activity-dependent synaptic modification.

Secreted molecules organize synapse differentiation

In addition to adhesion molecules, some secreted molecules also serve as synaptic organizers (Fig. 4). For example, the motor neuron–derived ligand agrin, which was the first identified secreted organizing molecule for postsynaptic differentiation, activates MuSK, a postsynaptic receptor tyrosine kinase, to regulate NMJ specialization (Glass et al., 1996; Zhou et al., 1999). Recently, a low-density lipoprotein receptor–related protein, LRP4, was identified as the co-receptor of agrin, forming a complex with MuSK and mediating MuSK signaling (Kim et al., 2008; Zhang et al., 2008). Several Wnts appears to act together with agrin to activate the LRP4–MuSK receptor complex to promote postsynaptic differentiation (Jing et al., 2009; Zhang et al., 2012). LRP4 also acts as a direct retrograde signal, functioning independently of MuSK for presynaptic differentiation (Yumoto et al., 2012), demonstrating that LRP4 acts as a bi-directional synaptic organizer (Fig. 4, left).Open in a separate windowFigure 4.Secreted trans-synaptic signaling at NMJs and CNS synapses. (Left) At Drosophila neuromuscular junctions (NMJs), Wnts are secreted from presynaptic terminals in association with Evi in the form of exosomes. In vertebrate NMJs, Wnt binds to the Agrin–LRP4–MuSK complex to regulate synapse formation. (Right) At CNS synapses, glia-derived thrombospondins (TSPs) and presynaptic neuron–derived cerebellin (Cbln) organize synapse differentiation and formation bi-directionally through binding to GluD2 and an isoform of neurexin (S4+) on the postsynaptic and presynaptic membranes, respectively. LTCC, L-type Ca²⁺ channel complex; AChR, acetyl choline receptor.Wnt is another well-characterized signaling molecule regulating many developmental processes including synaptic differentiation bi-directionally. Wnt regulates synaptic assembly both positively and negatively. For example, Wnt3 collaborates with agrin to promote the clustering of acetyl choline receptor (AChR) at the vertebrate NMJs (Henriquez et al., 2008), while Wnt3a inhibits AChR aggregation through β-catenin signaling (Wang et al., 2008). In the C. elegans NMJ, a Wnt molecule, CWN-2, stimulates the delivery and insertion of AchR to the postsynaptic membrane through the activation of a Frizzled–CAM-1 receptor complex (Jensen et al., 2012). Local Wnt gradient can suppress synapse formation in both C. elegans and Drosophila (Inaki et al., 2007; Klassen and Shen, 2007). Interestingly, in these contexts, Wnts are secreted from nonneuronal or nonsynaptic partner cells, suggesting that environmental factors can shape synaptic connections. Wnt can also be secreted from presynaptic neurons. A recent study demonstrated the trans-synaptic transmission of Wnt by exosome-like vesicles containing the Wnt-binding protein Evi at Drosophila NMJs (Fig. 4, left; Korkut et al., 2009; Koles et al., 2012). Presynaptic vesicular release of Evi is required for the secretion of Wnt. Intriguingly, different Wnt ligands regulate synapse formation in distinct cellular contexts. Wnt3a promotes excitatory synaptic assembly through CaMKII, whereas Wnt5a mediates inhibitory synapse formation by stabilizing GABA_A receptors (Cuitino et al., 2010; Ciani et al., 2011). This functional diversity indicates that different Wnts, receptors, and downstream pathways, as well as cell-specific contexts dictate the action of extracellular cues. Another conserved secreted molecule, netrin/UNC-6, can also pattern synapses by either promoting or inhibiting synapse formation (Colón-Ramos et al., 2007; Poon et al., 2008). Because Wnt and netrin often exist in gradients, these observations suggest that the localization of synapses can be specified by the gradient of extrinsic cues.In mammalian, several glia-derived cues have been shown to play important roles in regulating synapse formation or elimination. Thrombospondins (TSPs) are trans-synaptic organizers secreted from immature astrocytes (Christopherson et al., 2005). Both in vitro and in vivo data demonstrate the capacity of TSPs to increase synapse number, promote the localization of synaptic molecules, and refine the pre- and postsynaptic alignment (Christopherson et al., 2005; Eroglu et al., 2009). Recently, two transmembrane molecules were uncovered in mediating TSP-induced synaptogenesis (Fig. 4, right). Neuroligin 1 interacts with TSP1 with its extracellular domain mediating the acceleration of synaptogenesis in hippocampal neurons (Xu et al., 2010). α2δ-1, a subunit of the L-type Ca²⁺ channel complex (LTCC), was also identified as the postsynaptic receptor of TSP in excitatory CNS neurons (Eroglu et al., 2009). Interaction between TSP and α2δ–1 triggers the conformational changes and sequentially recruits synaptic scaffolding molecules and initiates synapse formation (Eroglu et al., 2009). Interestingly, TSP-induced synapses, although structurally normal and presynaptically active, are postsynaptically silent due to the lack of AMPA receptors (Christopherson et al., 2005), indicating the existence of other glia-derived signals involved in synapse formation. In fact, in cultured hippocampal neurons, a glia-derived neurotrophic factor GDNF enhances the pre- and postsynaptic adhesion by triggering the trans-homophilic interaction of its receptors GFRα1 localized at both pre- and postsynaptic sites (Ledda et al., 2007). Several other glia-derived factors have been shown to play critical roles in synaptogenesis. Astrocytes secrete extracellular molecules hevin and SPARC to regulate synapse formation in vitro and in vivo (Kucukdereli et al., 2011). Astrocytes also express a transmembrane adhesion protein, protocadherin-γ, serving as a local cue to promote synapse formation (Garrett and Weiner, 2009). TGF-β secreted from the NMJ glia acts together with the muscle-derived TGF-β to control synaptic growth (Fuentes-Medel et al., 2012). In a similar fashion, secretion of BDNF by vestibular supporting cells is required for synapse formation between hair cells and sensory organs (Gómez-Casati et al., 2010).Another important synaptic organizer is cerebellin (Cbln), a presynapse-derived complement protein, C1q-like family protein. In cbln1-null mice the number of parallel fibers (PF)–Purkinje synapses is dramatically reduced; the postsynaptic densities in the remaining synapses are larger than the apposite active zones (Hirai et al., 2005). Cbln was also found to regulate synaptic plasticity, as cbln1-null mice show impaired long-term depression in cerebellum (Hirai et al., 2005). These defects precisely resemble those in mice lacking a putative glutamate receptor, GluD2 (Kashiwabuchi et al., 1995; Kurihara et al., 1997), suggesting that Cbln1 and GluD2 function in synaptic differentiation through a common pathway. Interestingly, the C-terminal domain and N-terminal domains of GluD2 are indispensable for cerebella LTD and PF–Purkinje synaptic morphology, respectively (Kohda et al., 2007; Uemura et al., 2007; Kakegawa et al., 2008, 2009). Further studies suggested that Cbln1 directly binds to the N-terminal domain of GluD2 and recruits postsynaptic proteins by clustering GluD2 (Matsuda et al., 2010). Neurexin was recently reported as the presynaptic receptor of Cbln in promoting synaptogenesis (Uemura et al., 2010), which reinforces the understanding of Cbln-mediated trans-synaptic signaling: Cbln serves as a bi-directional synapse organizer by linking presynaptic neurexin and postsynaptic GluD2 (Fig. 4, right).Besides being required for synapse formation at early stages, genetic ablation of GluD2 in adult cerebellum leads to loss of PF–Purkinje synapses (Takeuchi et al., 2005), indicating that Cbln1–GluD2 signaling is also important for the maintenance of PF–Purkinje synapses. Chronic stimulation of neural activity decreases Cbln1 expression and diminishes the number of PF–Purkinje synapses (Iijima et al., 2009), suggesting the importance of Cbln1–GluD2 signaling for synaptic plasticity and homeostasis.Cbln subfamily proteins are widely expressed throughout the brain (Miura et al., 2006), suggesting that their synaptogenic roles may be wide spread in other regions of the brain. Cbln2 and 4 are also secreted proteins, whereas Cbln3 is retained in the cellular endomembrane system (Iijima et al., 2007). Cbln1 and 2, interacting with an isoform of presynaptic neurexin, induce synaptogenesis (Joo et al., 2011; Matsuda and Yuzaki, 2011). Notably, the cortical synapses induced by neurexin–Cbln signaling are preferentially inhibitory (Joo et al., 2011), distinguishing the effects of Cbln from neuroligin. GluD1 was recently found to be the postsynaptic receptor of Cbln1 and 2 in cortical neurons, mediating the differentiation of inhibitory presynapses (Yasumura et al., 2012). On the other side, Cbln4 selectively binds to the netrin receptor DCC in a netrin-displaceable manner (Fig. 4, right), suggesting a potential function of Cbln4 through DCC signaling pathway (Iijima et al., 2007). Intriguingly, C1q, although sharing similar structure with Cbln, serves an opposite role by regulating the synapse elimination: C1q released from retinal ganglion cells refines the retinogeniculate connections by eliminating unneeded synapses (Stevens et al., 2007).

Concluding remarks

Synapse development is regulated in multiple steps. Research over the last few years have uncovered many regulatory mechanisms on how trafficking of synaptic material is regulated and how scaffold proteins act with cytoskeleton networks and trans-synaptic signaling to instruct the synapse formation. Nevertheless, our understanding of the cellular and molecular mechanisms regulating synapse development is still incomplete. For example, how is the direction, speed, and amount of synaptic material being transported specified? How is a synapse’s size determined? How is synapse type and strength specified through adhesive and secreted trans-synaptic signaling? How do the redundant synapse-inducing pathways interact with each other? Given the rapidly emerging improvements of technologies, especially super-resolution microscopy and high-throughput genomics and proteomics, the synapse development field will likely rapidly evolve in the near future.     

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.     

Cytokinetic astralogy

Division plane specification in animal cells has long been presumed to involve direct contact between microtubules of the anaphase mitotic spindle and the cell cortex. In this issue, von Dassow et al. (von Dassow et al. 2009. J. Cell. Biol. doi:10.1083/jcb.200907090) challenge this assumption by showing that spindle microtubules can effectively position the division plane at a distance from the cell cortex.Cell division, or cytokinesis, is accomplished via constriction of an equatorially localized contractile ring composed of filamentous actin and myosin II (Rappaport, 1996). Accurate division plane specification is essential to properly partition the cytoplasm and permit each daughter cell to receive a single copy of the genome. To ensure this accuracy, microtubules of the mitotic spindle signal to the cell cortex upon anaphase onset and promote assembly of the contractile ring between the separating chromosomes. The precise mechanism by which microtubules position the contractile ring, however, remains elusive.Early models on the nature of the spindle-derived signal proposed that astral rays (later found to be microtubules) position the division plane by either locally promoting contractility at the cell equator or inhibiting contractility at the cell poles (Rappaport, 1996). Recent evidence, though, suggests that distinct microtubule populations within a single cell provide multiple signals to promote accurate division (Canman et al., 2003; Bringmann and Hyman, 2005; Chen et al., 2008; Foe and von Dassow, 2008; von Dassow, 2009).The anaphase mitotic spindle contains several subtypes of microtubules, each of which is likely to contribute to division plane specification. Although kinetochore microtubules drive chromosome segregation during anaphase, nonkinetochore microtubules extend and maintain close proximity with the assembling central spindle (Mastronarde et al., 1993). Central spindle microtubules are highly stable (Salmon et al., 1976) and organize into an antiparallel bundled array between the separating chromosomes (Mastronarde et al., 1993). Preventing central spindle assembly usually results in a complete failure in cytokinesis, and prevents division plane specification in many cell types (Glotzer, 2005). Astral microtubules, however, are highly dynamic and grow out circumferentially from the centrosomes toward the cell cortex. Increasing evidence suggests that the astral microtubule signal inhibits contractility (see below; Canman et al., 2000; Kurz et al., 2002; Lewellyn et al., 2009).Regardless of the mechanism of division plane specification via microtubules, nearly all current models depend on direct contact between microtubules of the mitotic spindle and the cell cortex. Most of these models were based on observations that at the time of division plane specification, astral microtubules contact the cell cortex in nearly all systems studied. Nonkinetochore and/or central spindle microtubules have also been proposed to deliver critical contractile signals to the cell equator (Murata-Hori and Wang, 2002; Canman et al., 2003; Somers and Saint, 2003; Verbrugghe and White, 2004; Lewellyn et al., 2009; Vale et al., 2009). Yet in many cell types (especially early embryos), central spindle microtubules are at some distance from the cell cortex during division plane specification. Despite this, signal delivery for both astral and central spindle microtubules was proposed to occur via direct transport along microtubules to the cell cortex. The study of von Dassow et al. in this issue, however, indicates that accurate division plane specification does not require any close microtubule/cortical contact and may occur via a diffusion-based mechanism (see also Salmon and Wolniak, 1990).By treating echinoderm and Xenopus embryos with controlled levels of trichostatin A (TSA), which destabilizes acetylated dynamic microtubules via inhibition of the tubulin deacetylase HDAC6 (Matsuyama et al., 2002), von Dassow et al. (2009) were able to preferentially prevent astral microtubule growth while leaving central spindle microtubules intact. TSA treatment did not block anaphase onset or central spindle assembly, but resulted in the complete disruption of all direct microtubule contact with the cell cortex. Nevertheless, TSA-treated cells were able to undergo cytokinesis successfully (Fig. 1 A). The lack of astral microtubules in TSA-treated cells was also recapitulated by double centrosome ablation, and again the cells were able to undergo cytokinesis (Fig. 1 B). In both experiments, cytokinesis occurred in a timely manner, but the contractile ring was broader than in control cells. Together, these data suggest that spindle microtubules are sufficient to provide a diffusible stimulatory signal capable of defining the cell division plane without any direct contact with the cell cortex (von Dassow et al., 2009).Open in a separate windowFigure 1.Testing models of division plane specification by targeting distinct microtubule populations. By selectively eliminating astral microtubules with either controlled TSA-treatment (A) or by double centrosome ablation (B), von Dassow et al. (2009) provide strong evidence that microtubule contact with the cell cortex is not essential for successful cytokinesis. When a single centrosome was ablated, the division plane was displaced away from the ablated aster (B); this suggests that astral microtubules provide an inhibitory signal. Further, anucleate cells would only complete cytokinesis if the intracentrosomal distance exceeded the distance from the centrosomes to the cell cortex (C).The authors noticed that cytokinesis occurred selectively at a position with reduced microtubule density in control cells; therefore, they explored the role of astral microtubules in division plane positioning. By selectively ablating one centrosome just before anaphase onset, von Dassow et al. (2009) were also able to provide strong support for an inhibitory role of astral microtubules in division plane specification. When a single centrosome was ablated, the division plane was displaced away from the remaining astral microtubules and toward the ablated centrosome (Fig. 1 B). Further evidence for an inhibitory role of astral microtubules in cytokinesis came from close examination of the intracentrosomal distance in anucleate cells that were able to undergo cytokinesis relative to those that were not. Cells were only able to undergo cytokinesis when the intracentrosomal distance exceeded the distance from the centrosomes to the cell cortex (Fig. 1 C), which suggests that cytokinesis requires an aster-free zone. The authors propose a mechanism in these anucleate cells whereby global activation of contractility drives division plane specification refined by a zone of astral separation (von Dassow et al., 2009). However, one possibility is that a central spindle still forms in these anucleate cells and thus provides the same diffusion-based signal that promotes division in cells without asters. Indeed, antiparallel arrays of bundled microtubules that resemble the central spindle are known to form between asters without intervening chromosomes in other systems (Savoian et al., 1999).To summarize, the results described by von Dassow et al. (2009) support a model in which central spindle microtubules provide a diffusible stimulatory signal to promote the assembly of a broad contractile ring, which is then refined by astral microtubules into a tight contractile ring. It is tempting to speculate on the molecular nature of this diffusible signal and mechanism of the astral refinement during cytokinesis. Signaling via the small GTPase Rho is required for cytokinesis and is dependent on spindle microtubules (Bement et al., 2005; Piekny et al., 2005). von Dassow et al. (2009) showed that in TSA-treated cells lacking astral microtubules, the equatorial zone of active Rho GTPase is broader relative to control cells. Rho activation is promoted (at least in part) via the central spindle–localized GTP exchange factor, ECT2 (Glotzer, 2005). In parallel, the GTPase-activating protein (GAP) CYK4/MgcRacGAP also associates with the central spindle, where it acts to both limit the zone of Rho activity (Miller and Bement, 2009) and to promote the inactivation of another small GTPase, Rac (D''Avino et al., 2004; Yoshizaki et al., 2004; Canman et al., 2008). Perhaps in parallel to central spindle mediated activation of Rho signaling, local inactivation of the inhibitory Rac signal via CYK4 GAP activity would further specify the division plane, even at a distance (Fig. 2). When the dynamic asters are present, they could then additionally amplify Rac signaling at the cell poles via a similar mechanism to what occurs during cell motility (Wittmann and Waterman-Storer, 2001). This local feedback loop would reinforce the positive signal coming from the central spindle via Rho activation and could help delimit active Rho at the cell equator (Fig. 2). Certainly, understanding how Rho activation can be propagated to the cell cortex via diffusion in such an accurate manner will be a major future challenge.Open in a separate windowFigure 2.Model for central spindle–mediated signaling via Rho family small GTPases. Central spindle–localized guanine nucleotide exchange factor ECT2 leads to Rho activation at the cell equator. At the same time, central spindle–localized CYK4 (a Rho family GAP) would also locally inactivate the inhibitory Rac signal. Further refinement of the zone of active Rho by astral microtubule–activated Rac could then sharpen the Rho zone into a tight contractile ring.     

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.