Interplay between Velocity and Travel Distance of Kinesin-based Transport in the Presence of Tau

Although the disease-relevant microtubule-associated protein tau is known to severely inhibit kinesin-based transport in vitro, the potential mechanisms for reversing this detrimental effect to maintain healthy transport in cells remain unknown. Here we report the unambiguous upregulation of multiple-kinesin travel distance despite the presence of tau, via decreased single-kinesin velocity. Interestingly, the presence of tau also modestly reduced cargo velocity in multiple-kinesin transport, and our stochastic simulations indicate that the tau-mediated reduction in single-kinesin travel underlies this observation. Taken together, our observations highlight a nontrivial interplay between velocity and travel distance for kinesin transport, and suggest that single-kinesin velocity is a promising experimental handle for tuning the effect of tau on multiple-kinesin travel distance.Conventional kinesin is a major microtubule-based molecular motor that enables long-range transport in living cells. Although traditionally investigated in the context of single-motor experiments, two or more kinesin motors are often linked together to transport the same cargo in vivo (1–4). Understanding the control and regulation of the group function of multiple kinesins has important implications for reversing failure modes of transport in a variety of human diseases, particularly neurodegenerative diseases. Tau is a disease-relevant protein enriched in neurons (5,6). The decoration of microtubules with tau is known to strongly inhibit kinesin transport in vitro (7–9), but how kinesin-based transport is maintained in the presence of high levels of tau, particularly in healthy neurons, remains an important open question. To date, no mechanism has been directly demonstrated to reverse the inhibitory effect of tau on kinesin-based transport. Here we present a simple in vitro study that demonstrates the significant upregulation of multiple-kinesin travel distance with decreasing ATP concentration, despite the presence of tau.This investigation was motivated by our recent finding that single-kinesin velocity is a key controller for multiple-kinesin travel distance along bare microtubules (10). The active stepping of each kinesin motor is stimulated by ATP (11), and each kinesin motor remains strongly bound to the microtubule between successive steps (10,11). As demonstrated for bare microtubules (10), with decreasing ATP concentrations, each microtubule-bound kinesin experiences a decreased stepping rate per unit time and spends an increased fraction of time in the strongly bound state; additional unbound kinesins on the same cargo have more time to bind to the microtubule before cargo travel terminates. Thus, reductions in single-kinesin velocity increase the probability that at least one kinesin motor will remain bound to the microtubule per unit time, thereby increasing the travel distance of each cargo (10). Because this effect only pertains to the stepping rate of each individual kinesin and does not address the potential presence of roadblocks such as tau on the microtubules, we hypothesized in this study that single-kinesin velocity may be exploited to relieve the impact of tau on multiple-kinesin travel distance.We focused our in vitro investigation on human tau 23 (htau23, or 3RS tau), an isoform of tau that exhibits the strongest inhibitory effect on kinesin-based transport (7–9). Importantly, htau23 does not alter the stepping rate of individual kinesins (7,9), supporting our hypothesis and enabling us to decouple single-kinesin velocity from the potential effects of tau. We carried out multiple-kinesin motility experiments using polystyrene beads as in vitro cargos (8,10), ATP concentration as an in vitro handle to controllably tune single-kinesin velocity (10,11), and three input kinesin concentrations to test the generality of potential findings for multiple-kinesin transport. Combined with previous two-kinesin studies (10,12), our measurements of travel distance (Fig. 1 A) indicate that the lowest kinesin concentration employed (0.8 nM) corresponds to an average of ∼2–3 kinesins per cargo. Note that in the absence of tau, the observed decrease in bead velocity at the higher kinesin concentrations (Fig. 1 A) is consistent with a recent in vitro finding (13). At 1 mM ATP, htau23 reduced kinesin-based travel distance by a factor of two or more (Fig. 1, A and B). This observation is in good agreement with previous reports (7,8).Open in a separate windowFigure 1Distributions of multiple-kinesin travel distances measured at three experimental conditions, to verify the effect of tau (A and B) and to investigate the impact of single-kinesin velocity on the tau effect (B and C). Shaded bars at 8.7 μm indicate counts of travel exceeding the field of view. The mean travel distance (d; ± standard error of mean, SEM), sample size (n), and corresponding mean velocity (v; ± SEM) are indicated. MT denotes microtubule. Mean travel distance increased substantially at 20 μM ATP (C), despite the presence of htau23. This effect persisted across all three kinesin concentrations tested (left to right).Consistent with our hypothesis, reducing the available ATP concentration to 20 μM increased the multiple-kinesin travel distance by >1.4-fold for all three input kinesin concentrations (Fig. 1, B and C), despite the presence of htau23. The corresponding reduction in single-kinesin velocity with decreasing ATP concentration (10,11) is reflected in the ∼3.4-fold reduction in the measured bead velocities (Fig. 1, B and C). Therefore, the strong negative relationship between single-kinesin velocity and multiple-kinesin travel distance occurs not only for bare microtubules (10), but also for tau-decorated microtubules.What causes the observed increase in travel distance at the lower ATP concentration (Fig. 1, B and C)? In addition to the mechanism discussed above for the case of bare microtubules (10), an intriguing mechanism was suggested by recent studies of tau-microtubule interactions in which htau23 was observed to dynamically diffuse along microtubule lattices (14,15): reducing the stepping rate of a microtubule-bound kinesin may effectively increase the probability that a tau roadblock can diffuse away before the kinesin takes its next step.Perhaps surprisingly, although htau23 does not impact single-kinesin velocity (7,9), we observed a modest reduction in the average velocity of multiple-kinesin transport in experiments using tau-decorated microtubules (Fig. 1, A and B). This decreased velocity reflects a substantially larger variance in the instantaneous velocity for bead trajectories in the presence of htau23 (see Fig. S1 in the Supporting Material), as quantified by parsing each bead trajectory into a series of constant-velocity segments using a previously developed automatic software incorporating Bayesian statistics (16).To test the possibility that single-kinesin travel distance impacts multiple-kinesin velocity, we performed stochastic simulations (see the Supporting Material) that assumed N identical kinesin motors available for transport and included kinesin’s detachment kinetics (17). Previously, this model successfully captured multiple-dynein travel distances in vivo using single-dynein characteristics measured in vitro (18). In this study, we introduced one (and only one) free parameter to reflect the probability of each bound kinesin encountering tau at each step. When encountering tau, each kinesin has a 54% probability of detaching from the microtubule (interpolated from Fig. 2A of Dixit et al. (7)); the undetached kinesin is assumed to remain engaged in transport and completes its step along the microtubule despite the presence of tau.Remarkably, our simple simulation suggested that the tau-mediated reduction in single-kinesin travel is sufficient to reduce multiple-kinesin velocity (Fig. 2 A). The majority of the velocity decrease is predicted to occur at the transition from single-kinesin to two-kinesin transport (Fig. 2). Further decreases in cargo velocity with increasing motor number are predicted to be modest and largely independent of tau (Fig. 2 B). The results of our simulation remain qualitatively the same when evaluated at two bounds (40 and 65%) encompassing the interpolated 54% probability of kinesin detaching at tau (see Fig. S2).Open in a separate windowFigure 2Stochastic simulations predict a tau-dependent reduction in multiple-kinesin velocity, assuming that the only effect of tau protein is to prematurely detach kinesin from the microtubule (or, to reduce single-kinesin travel distance). (A) Average velocity of cargos carried by the indicated number of kinesins was evaluated at 1 mM ATP, and for four probabilities that a kinesin may encounter tau at each step. Mean velocity was evaluated using 600 simulated trajectories for all simulation conditions. Error bars indicate SEM. (B) Change in cargo velocity with each additional kinesin (ΔVel/kinesin) as a function of tau-encounter probability. These values were calculated from cargo velocities shown in panel A. Error bars indicate SEM.We note that our simple simulations do not consider the possibility that kinesin may pause in front of a tau roadblock, as previously reported in Dixit et al. (7). We omitted this consideration because the interaction strength between kinesin and the microtubule in such a paused state is unknown. In a multiple-motor geometry, could a paused kinesin be dragged along by the other motors bound to the same cargo? Could a tau roadblock be forcefully swept off the microtubule surface by the collective motion of the cargo-motor complex? Significant experimental innovations are necessary to specifically address these questions in future multiple-motor assays and to guide modeling efforts. Nonetheless, our simple simulation demonstrates that reducing single-kinesin travel distance is sufficient to decrease multiple-kinesin travel distance.Taken together, our observations highlight a nontrivial interplay between velocity and travel distance for kinesin-based transport in the presence of tau. We uncover a previously unexplored dual inhibition of tau on kinesin-transport: in addition to limiting cargo travel distance, the tau-mediated reduction in single-kinesin travel distance also leads to a modest reduction in multiple-kinesin velocity. We provide what we believe to be the first demonstration of the unambiguous upregulation of multiple-kinesin travel distance despite the presence of tau, via reducing single-kinesin velocity, suggesting a mechanism that could be harnessed for future therapeutic interventions in diseases that result from aberrant kinesin-based transport.     

The contribution of αβ-tubulin curvature to microtubule dynamics

Microtubules are dynamic polymers of αβ-tubulin that form diverse cellular structures, such as the mitotic spindle for cell division, the backbone of neurons, and axonemes. To control the architecture of microtubule networks, microtubule-associated proteins (MAPs) and motor proteins regulate microtubule growth, shrinkage, and the transitions between these states. Recent evidence shows that many MAPs exert their effects by selectively binding to distinct conformations of polymerized or unpolymerized αβ-tubulin. The ability of αβ-tubulin to adopt distinct conformations contributes to the intrinsic polymerization dynamics of microtubules. αβ-Tubulin conformation is a fundamental property that MAPs monitor and control to build proper microtubule networks.Microtubules are polar polymers formed from αβ-tubulin heterodimers. These tubulin subunits associate head-to-tail to form protofilaments, and typically 13 protofilaments are associated side-by-side to form the hollow cylindrical microtubule. Most microtubules emanate from microtubule organizing centers, in which their minus ends are embedded. GTP-tubulin associates with the fast-growing plus ends as the microtubules radiate to explore the cell interior (see Box).

The cycle of microtubule polymerization.

Fig. 1). The addition of a new subunit completes the active site for GTP hydrolysis, and consequently most of the body of the microtubule contains GDP-bound αβ-tubulin. The GDP lattice is unstable but protected from depolymerization by a stabilizing “GTP cap,” an extended region of newly added GTP- or GDP.Pi-bound αβ-tubulin. The precise nature of the microtubule end structure and the size and composition of the cap are a matter of debate. Loss of the stabilizing cap leads to rapid depolymerization, which is characterized by an apparent peeling of protofilaments. “Catastrophe” denotes the switch from growth to shrinkage, and “rescue” denotes the switch from shrinkage to growth.Open in a separate windowFigure 1.Three structures of GTP-bound αβ-tubulin adopt similar curved conformations. Different αβ-tubulin structures were superimposed using α-tubulin as a reference, and oligomers were generated by assuming that the spatial relationship between α- and β-tubulin within a heterodimer is identical to the relationship between heterodimers. Curvature is calculated from the rotational component of the transformation required to superimpose the α-tubulin chain onto the β-tubulin chain of the same heterodimer. All of the GTP-bound structures (Rb3 complex, Protein Data Bank [PDB] accession no. 3RYH [magenta]; DARPin complex, PDB accession no. 4DRX [green]; TOG1 complex, PDB accession no. 4FFB [blue]) show between 10° and 13° of curvature, which is very similar to the curvature observed in GDP-bound structures (see inset, where the αβ-tubulins from a GDP-bound stathmin complex [PDB accession no. 1SA0] are shown in yellow and orange). A straight protofilament (putty and dark red color, PDB accession no. 1JFF) and a partially straightened assembly (tan) from GMPCPP ribbons are shown for reference.Unlike actin filaments, which grow steadily, microtubules frequently switch between phases of growth and shrinkage. This hallmark property of microtubules, known as “dynamic instability” (Mitchison and Kirschner, 1984), allows the microtubule cytoskeleton to be remodeled rapidly over the course of the cell cycle. “Catastrophes” are GTPase-dependent transitions from growing to shrinking, whereas “rescues” are transitions from shrinking to growing. Numerous microtubule-associated proteins (MAPs) regulate microtubule polymerization dynamics. Discovering how cells regulate and harness dynamic instability is a fundamental challenge in cell biology.A recent accumulation of structural, biochemical, and in vitro reconstitution data has advanced the understanding of dynamic instability and the MAPs that control it. Fresh structural data have provided insight into the process of microtubule assembly and defined how some MAPs recognize αβ-tubulin in and out of the microtubule. In vitro reconstitution experiments are reshaping the understanding of catastrophe and also providing quantitative insight into the mechanism of MAPs. Here, we review this progress, paying special attention to the emerging theme of interactions that are selective for different conformations of αβ-tubulin, both inside and outside the microtubule lattice. We argue for the central importance of recognizing these distinct conformations in the control of microtubule dynamics by MAPs and hence in the construction of a functional microtubule cytoskeleton by cells.

Tubulin dimers and their curvatures

It was clear in early EM studies that αβ-tubulin could form a diversity of polymers (Kirschner et al., 1974). In particular, the first cryo-EM of dynamic microtubules (Mandelkow et al., 1991) revealed significant differences in the appearance of growing and shrinking microtubule ends. Growing microtubule ends had straight protofilaments and were tapered, with uneven protofilament lengths, whereas shrinking microtubule ends had curved protofilaments that peeled outward and lost their lateral contacts. These and other data established the canonical model that GTP-tubulin is “straight” but GDP-tubulin is “curved” (Melki et al., 1989). The idea that GTP binding straightened αβ-tubulin into a microtubule-compatible conformation before polymerization was appealing because it provided a structural rationale for why microtubule assembly required GTP and how GTP hydrolysis could lead to catastrophe. A subsequent cryo-EM study (Chrétien et al., 1995), however, revealed that growing microtubules often tapered and curved gently outward without losing their lateral contacts. These data suggested that GTP-tubulin might not be fully straight at the time of its incorporation into the microtubule lattice, an observation that set the stage for a still-active debate on the structure of GTP-tubulin and of microtubule ends.The atomic details of “straight” and “curved” became apparent when the first structures of αβ-tubulin were solved. The straight conformation of αβ-tubulin was determined from cryo-electron crystallographic studies of Zn-induced αβ-tubulin sheets (Nogales et al., 1998). The structure showed linear head-to-tail stacking of αβ-tubulin along the protofilament, both within and between αβ-tubulin heterodimers. The curved conformation of αβ-tubulin was determined from x-ray crystallographic studies of a complex between αβ-tubulin and Rb3 (Gigant et al., 2000; Ravelli et al., 2004), a microtubule-destabilizing factor in the Op18/stathmin family (Belmont and Mitchison, 1996). In this complex, the individual α- and β-tubulin chains adopted a characteristic conformation distinct from their straight one. Longitudinal interactions also differed from those in the straight conformation (Fig. 1): within and between the heterodimers, successive α- and β-tubulin chains were related by an ∼12° rotation. A chain of these curved αβ-tubulins generates an arc with a radius of curvature resembling that of the peeling protofilaments at shrinking microtubule ends (Gigant et al., 2000; Steinmetz et al., 2000).Straight and curved are not the only two conformations, however. A cryo-EM study of αβ-tubulin helical ribbons trapped using guanylyl 5′-α,β-methylenediphosphonate (GMPCPP), a slowly hydrolyzable analogue of GTP, provided a molecular view of a possible microtubule assembly intermediate (Wang and Nogales, 2005). In these ribbons, GMPCPP-bound αβ-tubulin adopted a conformation roughly halfway (∼5° rotation) between the straight and curved conformations. These partially curved αβ-tubulin heterodimers formed two types of lateral bonds, only one of which resembled those in the microtubule. This structure suggested that at least some αβ-tubulin straightening occurs during polymerization.Until recently, structural information about the conformation of unpolymerized GTP-bound αβ-tubulin was notably lacking. Three recent crystal structures (Nawrotek et al., 2011; Ayaz et al., 2012; Pecqueur et al., 2012) have now provided remarkably similar views of this previously elusive species. In all three structures, GTP-bound αβ-tubulin adopts a fully curved conformation, with its α- and β-tubulin subunits related by ∼12° of rotation (Fig. 1). This curvature is not consistent with models in which GTP binding straightens unpolymerized αβ-tubulin. In each of the structures, αβ-tubulin is bound to another protein, stathmin/Rb3 (Ozon et al., 1997), a designed ankyrin repeat protein (DARPin; Pecqueur et al., 2012), as well as a TOG domain from the Stu2/XMAP215 family of microtubule polymerases (Gard and Kirschner, 1987; Wang and Huffaker, 1997). Biochemical experiments have failed to detect GTP-induced straightening of αβ-tubulin, arguing against the possibility that these unrelated binding partners forced GTP-tubulin to adopt the curved conformation. For example, the affinity of stathmin–tubulin interactions is the same for GTP-tubulin and GDP-tubulin (Honnappa et al., 2003). Similarly, five small molecule ligands that target the colchicine binding site and are predicted to bind only curved αβ-tubulin have equivalent affinity for GTP-tubulin, GDP-tubulin, and αβ-tubulin in the stathmin complex (Barbier et al., 2010). Likewise, a TOG domain from Stu2p binds to GTP- and GDP-tubulin with comparable affinity (Ayaz et al., 2012). Finally, DARPin binds equally well to GTP- and GDP-tubulin even though it contacts a structural element that is positioned differently in the straight and curved conformations (Pecqueur et al., 2012). Taken together with early biochemical experiments (Manuel Andreu et al., 1989; Shearwin et al., 1994), these new data strongly support a model in which unpolymerized αβ-tubulin is curved whether it is bound to GTP or to GDP (Buey et al., 2006; Rice et al., 2008; Nawrotek et al., 2011). According to this model, the curved-to-straight transition occurs during the polymerization process, not before. We discuss some implications of this new view at the end of the following section.

Conformation and dynamic instability

How does GTP hydrolysis destabilize the microtubule lattice and trigger catastrophe? A recent structural study has compared high-resolution cryo-EM reconstructions of GMPCPP microtubules and GDP microtubules to provide some answers to this question (Alushin et al., 2014). The structures show that GTP hydrolysis induces a compaction at the longitudinal interface between dimers, immediately above the exchangeable nucleotide-binding site. This compaction is accompanied by conformational changes in α-tubulin. In contrast, lateral contacts between tubulins were essentially unchanged in the different nucleotide states. These observations suggest that GTP hydrolysis introduces strain into the lattice, but how this strain affects the strength of longitudinal and lateral bonds to destabilize the microtubule remains unknown. The GMPCPP and GDP microtubules also show distinct arrangements of elements that bind to MAPs, which suggests a structural mechanism some MAPs could use to distinguish GTP lattices from GDP lattices (discussed later).In parallel with these structural advances, in vitro reconstitutions (Gardner et al., 2011b) have undermined the textbook view about the kinetics of catastrophe. The seminal measurements of catastrophe frequency (Walker et al., 1988, 1991) assumed that catastrophe occurred with the same probability on newly formed and old microtubules. In other words, the analysis implied that catastrophe was a first-order, single-step process. Although subsequent experiments (e.g., Odde et al., 1995; Janson et al., 2003) indicated that catastrophe involved multiple steps, the first-order view of catastrophe was widely adopted (Howard, 2001; Phillips et al., 2008). Recent experiments using a single-molecule assay for microtubule growth (Gell et al., 2010) have now shown definitively that catastrophe is not a single-step process; rather, newly formed microtubules undergo catastrophe less frequently than older ones (Gardner et al., 2011b). “Age-dependent” catastrophe implies that the stabilizing structure at the end of growing microtubules is evolving to become less effective. The timescale of this evolution is long compared with the kinetics of αβ-tubulin association (Gardner et al., 2011a). Thus, the ageing process probably reports on one or more structural properties of the microtubule end, such as the presence of “defects” in the lattice (Gardner et al., 2011b) or possibly increased tapering of microtubule ends (Coombes et al., 2013).It now seems clear that changes in the curvature of αβ-tubulin during microtubule polymerization are fundamental to microtubule dynamics and the regulatory activities of MAPs. Having straight conformations of αβ-tubulin only occur appreciably in the microtubule lattice provides a simple structural mechanism by which MAPs can discriminate unpolymerized from polymerized αβ-tubulin. Biochemical properties that define microtubule dynamics, like the strength of lateral and longitudinal contacts and the rate of GTP hydrolysis, may differ for curved, straight, and intermediate conformations of αβ-tubulin; e.g., curved forms probably bind microtubule ends less tightly than straight forms. By regulating when and where these different conformations occur, MAPs can tune microtubule dynamics. More speculatively, the complex biochemistry associated with different conformations of αβ-tubulin may contribute to the aging of microtubule ends, which leads to catastrophe. Understanding the connections between αβ-tubulin conformation, biochemistry, and polymerization dynamics is a major challenge for the future. Expanding the current mathematical models (Bowne-Anderson et al., 2013) and computational models (VanBuren et al., 2005; Margolin et al., 2012) of microtubule dynamics to incorporate these new findings about αβ-tubulin structure and age-dependent catastrophe may yield significant insights. In the following sections, we will examine recent studies that demonstrate how MAPs use selective interactions with distinct conformations of αβ-tubulin to control microtubule dynamics and thereby the physiology of the microtubule cytoskeleton.

Microtubule depolymerases stabilize curved conformations of tubulin

Perhaps the first direct evidence that MAPs might control the conformation of αβ-tubulin came from studies of microtubule depolymerases, which are proteins that promote, accelerate, or induce the depolymerization of microtubules (Howard and Hyman, 2007). Cells use microtubule depolymerases to maintain local control of microtubule catastrophe. Early electron microscopy studies of two unrelated depolymerases, Op18/stathmin and the kinesin-13 Xkcm1, showed that these proteins were able to induce/stabilize the curved conformation of αβ-tubulin and/or curved protofilaments (Desai et al., 1999; Gigant et al., 2000; Steinmetz et al., 2000). Depolymerases are also referred to as “catastrophe factors” because they trigger catastrophes in dynamic microtubules. The localized control of catastrophe is the essential function of depolymerases in cell physiology.The microtubule depolymerase stathmin is inactivated around chromosomes and at the leading edge of migrating cells (Niethammer et al., 2004), creating a gradient of depolymerase activity in these zones. Proteins in the Op18/stathmin family form a tight complex with two curved tubulin dimers (Fig. 2 A). Op18/stathmin proteins have been critical for the crystallization of tubulin (Ravelli et al., 2004; Gigant et al., 2005; Prota et al., 2013) and for biochemical studies of tubulin conformation. Although stathmins are frequently described as tubulin-sequestering proteins, the effect they have on microtubule catastrophe frequencies in vitro is much stronger than would be predicted from the simple sequestration of tubulin (Belmont and Mitchison, 1996). The potency of stathmins suggests that they induce catastrophes through direct interactions with microtubule ends, presumably weakening the bonds of terminal subunits by inducing or stabilizing their curvature (Gupta et al., 2013).Open in a separate windowFigure 2.Proteins that recognize curved αβ-tubulin tend to make long interfaces that span both α- and β-tubulin. (A) A stathmin family protein (blue) forms a long helix that binds two αβ-tubulin heterodimers (pink and green; PDB accession no. 3RYH). (B) The structure of a complex between kinesin-1 and αβ-tubulin (PDB accession no. 4HNA) is shown with the motor in dark green and αβ-tubulin in pink and lime. Depolymerizing kinesins have insertions (red segments modeled based on a crystal structure of MCAK; PDB accession no. 1V8K), such as the KVD finger, that expand the contact region compared with purely motile kinesins. (C) The TOG1 domain (blue) from Stu2, an XMAP215 family polymerase, contacts regions of α- and β-tubulin (pink and green) that move relative to each other in the curved (left, PDB accession no. 4FFB) and straight (right, model substituting straight αβ-tubulin; PDB accession no. 1JFF) conformations of αβ-tubulin. The asterisks show where this relative movement would disrupt the TOG–tubulin interface. Red side chains indicate conserved tubulin-binding residues at the top and bottom of the TOG domain. (D) The TOG2 domain from human CLASP1 (light blue, PDB accession no. 4K92) shows an “arched” interface that in docked models like the ones shown here is not complementary to curved (left) or straight (right) conformations of αβ-tubulin. Curved and straight structures are PDB 4FFB and 1JFF, respectively. Red side chains indicate binding residues similar to those in the polymerase family TOG domains, and asterisks highlight where the arched nature of this TOG prevents a conserved binding residue from contacting its interaction partner on β-tubulin.Kinesin-13s, first identified by their central motor domain (Aizawa et al., 1992; Wordeman and Mitchison, 1995), depolymerize microtubules catalytically using the energy of ATP hydrolysis (Hunter et al., 2003). Kinesin-13s depolymerize microtubules at spindle poles to generate poleward flux (Ganem et al., 2005), at kinetochores to drive anaphase chromosome segregation (Maney et al., 1998; Rogers et al., 2004), and in neuronal processes (Homma et al., 2003). Evidence that kinesin-13s depolymerized microtubules came from the discovery of the Xenopus laevis homologue, Xkcm1, in a screen for kinesin-related proteins involved in spindle assembly (Walczak et al., 1996). Incubation of Xkcm1, also known as MCAK, with GMPCPP microtubules caused peeled protofilaments and significant “ram’s horns” structures to appear at microtubule ends (Desai et al., 1999), which indicates that MCAK binds more tightly to curved structures than to straight ones. As with all kinesins, tight binding of the motor domain is coupled to its ATP hydrolysis cycle. Kinesin-13s first bind the microtubule lattice with an on-rate constant that strongly influences its depolymerase activity (Cooper et al., 2010). Kinesin-13s then target the end of the microtubule via “lattice diffusion,” a random walk mediated by electrostatic interactions that occurs in the ADP state (Helenius et al., 2006). Exchange of ADP to ATP occurs at microtubule ends; in the ATP state, MCAK binds tightly to tubulin dimers and either induces or stabilizes their outward curvature and detachment from the microtubule lattice (Friel and Howard, 2011). The subsequent hydrolysis of ATP causes kinesin-13 to release its tubulin subunit, now detached from the lattice, and begin another cycle of depolymerization (Moores et al., 2002).A distinguishing feature of the kinesin-13 motor domain is an extension of loop L2, known as the KVD finger (Ogawa et al., 2004; Shipley et al., 2004), which protrudes from the motor domain toward the minus end of the microtubule (Fig. 2 B). Alanine substitution of the KVD motif inhibits depolymerase activity in cell-based assays (Ogawa et al., 2004) and in vitro (Shipley et al., 2004). A recent cryo-EM study showed that the kinesin-13 motor domain contacts curved tubulin on three distinct surfaces (Asenjo et al., 2013) that differ from the contact surfaces of kinesin-1 (Sindelar and Downing, 2010; Gigant et al., 2013). The location of the kinesin-13 contact surfaces could allow kinesin-13 to stabilize spontaneous curvature of tubulin dimers at either microtubule end. Alternatively, tight binding of the kinesin-13 motor domain could directly induce curvature in the tubulin dimer. In either case, by promoting curvature at the growing microtubule end, kinesin-13s weaken the association of terminal subunits and induce catastrophes.Kinesin-8s are motile depolymerases (Gupta et al., 2006; Varga et al., 2006) that establish the length of microtubules in the mitotic spindle (Goshima et al., 2005; Rizk et al., 2014), position the spindle (Gupta et al., 2006), and modulate the dynamics of kinetochore microtubules (Stumpff et al., 2008; Du et al., 2010). Unlike the nonmotile kinesin-13s, whose motor domain is fully specialized for depolymerization, kinesin-8 proteins walk to the microtubule end and remove tubulin upon arrival (Gupta et al., 2006; Varga et al., 2006). Although it is unclear if depolymerase activity is fully conserved (Du et al., 2010; Mayr et al., 2011), all kinesin-8s combine motility with a negative effect on microtubule growth. For Saccharomyces cerevisiae Kip3p, the combination of motility and depolymerase activity has a significant functional consequence: Kip3p depolymerizes longer microtubules faster than shorter ones (Varga et al., 2006). This length-dependent depolymerization can be explained by an “antenna model.” In this model, longer microtubules will accumulate more kinesin-8s, which then walk toward the microtubule end, forming length-dependent traffic jams in some cases (Leduc et al., 2012). Because the rate of depolymerization depends on the number of kinesin-8s that arrive at the microtubule end, longer microtubules will be depolymerized more quickly. The “antenna model” depends critically on the high processivity of kinesin-8, which is thought to result from an additional C-terminal microtubule-binding element (Mayr et al., 2011; Stumpff et al., 2011; Su et al., 2011; Weaver et al., 2011); the C terminus may also contribute to a recently described microtubule sliding activity in Kip3p (Su et al., 2013). Intriguingly, a single Kip3p appears to be insufficient to remove a tubulin dimer. Rather, a second Kip3p must arrive at the microtubule end to bump off the first one (Varga et al., 2009).There are less structural and mutagenesis data available to explain the unique ability of kinesin-8s to walk and depolymerize. It is also not clear that all kinesin-8s use the same cooperative mechanism described for Kip3p. Like kinesin-13, the motor domain of kinesin-8 has an extended loop L2. This loop is disordered in the available crystal structure, but has been observed to contact α-tubulin in a cryo-EM reconstruction (Peters et al., 2010). The kinesin-8 loop L2 lacks a KVD sequence, however, and systematic mutations of L2 have not yet determined its role in depolymerase activity. The extent to which kinesin-8s recognize/induce curvature at microtubule ends remains unresolved. Truncated kinesin-8 motor domains can create small peels at the ends of GMPCPP microtubules (Peters et al., 2010), which suggests that kinesin-8 can induce or stabilize curvature. The fact that two kinesin-8s are required to dissociate a tubulin subunit, however, indicates that single motors alone do not substantially weaken the bonds holding the terminal tubulin subunit. Perhaps kinesin-8s do not stabilize curved forms of αβ-tubulin as strongly as kinesin-13s do.Reconstitution of microtubule dynamics in vitro showed that the depolymerizing kinesins affect catastrophe in different ways (Gardner et al., 2011b): kinesin-13s eliminate the aging process described earlier, whereas kinesin-8s accelerate it. Importantly, the local control of catastrophes by depolymerases is accomplished primarily through the local modulation of curvature at microtubule ends.

Growth-promoting MAPs also use conformation-selective interactions with αβ-tubulin

MAPs that accelerate growth or stabilize the microtubule lattice counteract microtubule depolymerases (Tournebize et al., 2000; Kinoshita et al., 2001). XMAP215 was discovered as the major protein in Xenopus extracts that promotes microtubule growth (Gard and Kirschner, 1987). Later, functional homologues were discovered in S. cerevisiae (Stu2p) (Wang and Huffaker, 1997) and other organisms (e.g., Charrasse et al., 1998; Cullen et al., 1999). XMAP215 family proteins localize to kinetochores and microtubule organizing centers, where they contribute to chromosome movements and to spindle assembly and flux (Wang and Huffaker, 1997; Cullen et al., 1999). Loss of XMAP215 family polymerase function leads to shorter, slower-growing microtubules and often gives rise to smaller and/or aberrant spindles (Wang and Huffaker, 1997; Cullen et al., 1999). All family members contain multiple TOG domains that bind αβ-tubulin (Al-Bassam et al., 2006; Slep and Vale, 2007). The molecular mechanisms underlying the activity of these proteins, and the collective action of their arrayed TOG domains, have until recently remained obscure. Recent progress is defining the structure and biochemistry of TOG domains and their interactions with αβ-tubulin. The emerging view is that XMAP215 family polymerases, like the depolymerases, bind to curved αβ-tubulin dimers as an important part of their biochemical cycle. In this section, we will focus on the most recent developments that are shaping the molecular understanding of growth-promoting MAPs, emphasizing the somewhat better studied XMAP215 family.Affinity chromatography using immobilized TOG domains from Stu2p revealed that the TOG1 domain binds directly to unpolymerized αβ-tubulin (Al-Bassam et al., 2006). TOG domains can also bind specifically to one end of the microtubule (Al-Bassam et al., 2006). Crystal structures of TOG domains, sequence conservation, and site-directed mutagenesis defined the αβ-tubulin–interacting surface, which forms a narrow “spine” of the book-shaped domain (Al-Bassam et al., 2007; Slep and Vale, 2007).In early models for XMAP215, the arrayed TOG domains were thought to bind multiple αβ-tubulins (Gard and Kirschner, 1987). Subsequent fluorescence-based reconstitution of XMAP215 activity, however, gave results that were not consistent with this “shuttle” model (Brouhard et al., 2008). The reconstitution assays showed that XMAP215 acted processively, residing at the microtubule end long enough to perform multiple rounds of αβ-tubulin addition. Intriguingly, XMAP215 increased the rate of, but not the apparent equilibrium constant for, microtubule elongation. XMAP215 also stimulated the rate of shrinkage in the absence of unpolymerized αβ-tubulin. Similar observations were made using Alp14 (Al-Bassam et al., 2012), a Schizosaccharomyces pombe XMAP215 homologue. These studies showed that XMAP215 catalyzes polymerization: it promotes microtubule growth by using its TOG domains to repeatedly bind and stabilize an intermediate state that otherwise limits the rate of polymerization.How do TOG domains recognize the microtubule end and promote elongation? Recent structural studies (Ayaz et al., 2012, 2014) suggest that interactions with curved αβ-tubulin play a central role. The crystal structures of complexes between αβ-tubulin and the TOG1 or TOG2 domains from Stu2p revealed that both TOG domains bind to curved αβ-tubulin (Ayaz et al., 2012, 2014; Fig. 2 C). The TOG domains do not interact strongly with microtubules even though the TOG-contacting epitopes are accessible on the microtubule surface (Ayaz et al., 2012). Preferential binding to curved αβ-tubulin (Ayaz et al., 2014) occurs because the arrangement of the TOG-contacting regions of α- and β-tubulin differs between curved and straight conformations (Fig. 2 C). Conformation-selective TOG–αβ-tubulin interactions explain how XMAP215 family proteins discriminate unpolymerized αβ-tubulin from αβ-tubulin in the body of the microtubule. XMAP215 family proteins require a basic region in addition to TOG domains for microtubule plus end association and polymerase activity (Widlund et al., 2011). The polarity of TOG–αβ-tubulin interactions and the ordering of domains in the protein together explain the plus end specificity of these polymerases: only at the plus end can TOGs engage curved αβ-tubulin while the C-terminal basic region contacts surfaces deeper in the microtubule (Ayaz et al., 2012). A recent study proposed that the linked TOG domains catalyze elongation using a tethering mechanism that effectively concentrates unpolymerized αβ-tubulin near curved subunits already bound at the microtubule end (Ayaz et al., 2014). The mechanisms by which these proteins catalyze depolymerization are less understood, although depolymerization can be explained by the catalytic stabilization of an intermediate state (Brouhard et al., 2008). By analogy with the depolymerases described earlier, the stabilization of such a state by arrayed TOG domains seems likely to also depend on the preferential interactions with curved αβ-tubulin.CLASP family proteins (Pasqualone and Huffaker, 1994; Akhmanova et al., 2001) also contain TOG domains, but they are used to different effect: CLASPs do not make microtubules grow faster but instead appear to regulate the frequencies of catastrophe and rescue. For example, in vitro reconstitutions using Cls1p, a CLASP protein from S. pombe, showed that Cls1p promoted rescue (Al-Bassam et al., 2010). CLASP family proteins also localize to kinetochores and contribute to spindle flux (Maiato et al., 2005). Loss of CLASP function affects microtubule stability and causes spindle defects (Akhmanova et al., 2001; Maiato et al., 2005), but does so without significantly affecting microtubule growth rates (Mimori-Kiyosue et al., 2006). CLASPs can also stabilize microtubule bundles/overlaps (Bratman and Chang, 2007). The recently published structure of a CLASP family TOG domain (Leano et al., 2013) provided an unexpected hint about a possible origin of the different activities. Indeed, the structure revealed significant differences with XMAP215 family TOG domains even though the CLASP TOG maintains evolutionarily conserved αβ-tubulin–interacting residues (Fig. 2 D). Whereas the αβ-tubulin binding surface of XMAP215 family TOGs is relatively flat, the equivalent surface of the CLASP TOG is arched in a way that appears to break the geometric match with curved αβ-tubulin (Leano et al., 2013; Fig. 2 D). This suggests that CLASP TOG domains might bind to an even more curved conformation of αβ-tubulin that has not yet been observed, that they do not simultaneously engage α- and β-tubulin, or that they do something else. It is not yet clear how these different possibilities might contribute to the rescue-promoting activity of CLASPs. However, even though the biochemical and structural understanding of how CLASP TOGs interact with αβ-tubulin is less advanced than for XMAP215 family TOGs, the conservation of critical αβ-tubulin–interacting residues makes it seem likely that conformation-selective interactions with αβ-tubulin will play a prominent role.The modulation of microtubule dynamics by XMAP215/CLASP family proteins ensures proper microtubule function in both interphase and dividing cells. As for the depolymerases, specific interactions with curved αβ-tubulin likely underlie the different regulatory activities of XMAP215/CLASP family proteins.

Sensing conformation at lattice contacts

Thus far, we have described how microtubule polymerases and depolymerases bind selectively to curved conformations of the αβ-tubulin dimer. These interactions play a significant role in the movement of tubulin dimers into and out of the microtubule polymer. Once in the polymer, αβ-tubulin dimers make contacts with neighboring tubulins. Recently, three MAPs were shown to bind microtubules at lattice contacts: (1) the Ndc80 complex, a core kinetochore protein; (2) doublecortin (DCX), a neuronal MAP; and (3) EB1, the canonical end-binding protein. Here we will summarize recent progress demonstrating how these proteins recognize distinctive features of lattice contacts.The Ndc80 complex is a core component of the kinetochore–microtubule interface (Janke et al., 2001; Wigge and Kilmartin, 2001; McCleland et al., 2003), forming a “sleeve” that connects the outer kinetochore to microtubules of the mitotic spindle (Cheeseman et al., 2006; DeLuca et al., 2006). Loss of Ndc80 function leads to chromosome segregation errors in mitosis (McCleland et al., 2004; DeLuca et al., 2005). Ndc80 binds to microtubules at the longitudinal interface between α- and β-tubulin and extends outward toward the plus end at an ∼60° angle (Cheeseman et al., 2006; Wilson-Kubalek et al., 2008). Ndc80 binds to both the intradimer and interdimer interface and forms oligomeric arrays (Alushin et al., 2010). The binding of Ndc80 to this longitudinal lattice contact may confer a preference for straight rather than curved microtubule lattices, because the shape of the Ndc80 binding site is expected to change as a protofilament bends (Alushin et al., 2010; Fig. 3 A). Preferential binding to straight protofilaments might allow the Ndc80 complex to remain attached to the end of a shrinking microtubule. Indeed, reconstitutions of the Ndc80 complex interacting with dynamic microtubules show that the curved shrinking end acts as a “reflecting wall,” giving rise to “biased diffusion” (Powers et al., 2009). Interestingly, the Ndc80 complex also promotes rescue (Umbreit et al., 2012), and selective binding to straight lattice contacts may contribute to this rescue activity.Open in a separate windowFigure 3.Proteins that bind microtubules can distinguish unique configurations at lattice contacts. (A) Ndc80 (light and dark blue) binds the contact within (dark blue) and between (light blue) αβ-tubulin heterodimers (pink and green). The left shows part of an Ndc80 array on straight protofilaments (PDB accession no. 3IZ0). The right shows that neighboring Ndc80 molecules clash when modeled onto a curved protofilament. Individual Ndc80s may read the conformation at a single joint, or the change in conformation may disrupt cooperative interactions between adjacent Ndc80s. (B) Two views of DCX (blue) binding a lattice contact at the vertex of four αβ-tubulins, PDB accession no. 4ATU. Cooperative interactions on the microtubule allow DCX to discriminate between the subtle changes that accompany different protofilament numbers (11: orange, EMDataBank [EMD] accession no. 5191; 13: red, EMD accession no. 5193; 15: yellow, EMD accession no. 5195). (C) EB1 (left, dark blue) binds at the same vertex as DCX (PDB accession no. 4AB0), but EB1 binds preferentially to GTP vertices over GDP vertices, and is not sensitive to protofilament number. The same section of microtubule with EB1 removed (right) shows the location of nucleotide-dependent changes at the four-way vertex: helix H3 of β-tubulin (red patch at the lower right of the four-way junction), and the intermediate (Int.) domain of α-tubulin (yellow patch at the top left of the four-way junction). pfs, protofilaments.DCX, a MAP expressed in developing neurons (Francis et al., 1999; Gleeson et al., 1999) and mutated in cases of subcortical band heterotopia (des Portes et al., 1998; Gleeson et al., 1998), is unique in its ability to bind specifically to 13-protofilament microtubules over other protofilament numbers (Moores et al., 2004; Fig. 3 B). DCX contains two nonidentical, microtubule-binding “DC” domains (Taylor et al., 2000) that share a ubiquitin-like fold (Kim et al., 2003). A cryo-EM reconstruction showed that a single DC domain binds to microtubules at the vertex of four tubulin dimers in the so-called “B” lattice configuration (Fourniol et al., 2010). The DCX binding site is ideally situated to detect the subtle changes at lattice contacts that result from different protofilament numbers, which range from 11 to 16 for mammalian microtubules (Sui and Downing, 2010). Despite their ideal location, protofilament preference is not a property of single DCX molecules. Rather, it is cooperative interactions between neighboring DCX molecules that are sensitive to the spacing between protofilaments (Bechstedt and Brouhard, 2012). In vitro, this selectivity enables DCX to nucleate homogeneous, 13-protofilament microtubules (Moores et al., 2004). The function of DCX in developing neurons remains unclear, with models ranging from microtubule stabilization (Gleeson et al., 1999) to regulation of kinesin traffic (Liu et al., 2012).EB1, the canonical end-binding protein (Morrison et al., 1998), uses its calponin homology (CH) domain (Hayashi and Ikura, 2003) to bind the same lattice contact as DCX (Maurer et al., 2012). EB1 forms “comets” by binding rapidly and tightly to a distinct feature at the growing microtubule end but only weakly to the “mature” lattice (Bieling et al., 2007). Recent work has defined this distinctive feature as the nucleotide state. EB1 binds preferentially to microtubules built from GTP analogues (Zanic et al., 2009; Maurer et al., 2011). Combined with careful analysis of the size, shape, and dynamics of EB1 comets (Bieling et al., 2007), these results established that EB1 recognizes microtubule ends by binding specifically to the “GTP cap,” which is an extended region of the microtubule end that is enriched with GTP- and GDP-Pi-tubulin dimers. A recent cryo-EM reconstruction of the CH domain of Mal3 (the S. pombe EB1) bound to GTPγS microtubules provided a possible structural mechanism for how EB1 might differentiate GTP from GDP lattices (Maurer et al., 2012; Fig. 3 C). Mal3 was observed to contact helix H3 of β-tubulin, which connects directly to the exchangeable nucleotide-binding site. EB1 also contacts the regions of α-tubulin that move during the compaction of the lattice that follows GTP hydrolysis (Alushin et al., 2014). Mutation of conserved EB1 residues that contact either helix H3 or the compacting region of α-tubulin disrupts the end-tracking behavior of EB1 (Slep and Vale, 2007; Maurer et al., 2012). Interactions with helix H3 and the compacting region of α-tubulin also enable EB1 to accelerate the transitions of tubulin from the GTP state to the GDP state; in other words, EB1 acts as a “maturation factor” for the microtubule end (Maurer et al., 2014). EB1 recruits a large network of plus-end-tracking proteins (Akhmanova and Steinmetz, 2008) through interactions with the EB1 C terminus (Hayashi et al., 2005; Honnappa et al., 2006) and EB1 homology domain (Honnappa et al., 2009). This diverse and complex protein network is essential for the regulation of microtubule dynamics, the capture of microtubule ends by the cell cortex (Kodama et al., 2003) and endoplasmic reticulum (Grigoriev et al., 2008), and the positioning of the mitotic spindle (Liakopoulos et al., 2003).As mentioned earlier, microtubule ends also show unique structural configurations, namely tapered, outwardly flared, and flattened structures collectively described as “sheets” (Chrétien et al., 1995). The sheets contain distinctive lattice contacts, and recent work shows that the microtubule-binding activities of DCX and EB1 are sensitive to these structural features. DCX, for example, binds specifically to the outwardly flared sheets (Bechstedt et al., 2014), which enables DCX to track microtubule ends. Evidence for the ability of EB1 to recognize or control a distinct lattice configuration comes from the reconstitutions showing that EB1 promotes elongation synergistically with XMAP215 (Zanic et al., 2013): lack of a detectable direct EB1–XMAP215 interaction suggested that the observed synergy was mediated through alterations of the microtubule end structure itself. Further evidence that EB1 can affect the structure of the microtubule lattice comes from data showing that EB1 can nucleate “A” lattice microtubules in vitro (des Georges et al., 2008) and influence protofilament number distributions (Vitre et al., 2008; Maurer et al., 2012). The connection between the structure of microtubule ends, their nucleotide state, and microtubule dynamics is an important open question.

Conclusions and outlook

The αβ-tubulin dimer adopts a range of conformations as it moves in and out of the microtubule polymer, including changes to its intrinsic curvature and changes to its lattice contacts. These different conformations affect microtubule dynamics by altering the strength of lattice association and the rate of GTP hydrolysis. The work we discussed here has revealed an intimate linkage between these different conformations and the activities of key proteins that regulate microtubule dynamics. It is now clear that selective interactions with distinct conformations of unpolymerized and polymerized αβ-tubulin define the cell physiology of the microtubule cytoskeleton. Recently developed methods for purifying or overexpressing αβ-tubulin (des Georges et al., 2008; Johnson et al., 2011; Widlund et al., 2012; Minoura et al., 2013) are facilitating structural studies and allowing the biochemistry of αβ-tubulin polymerization to be dissected in unprecedented detail. Microtubule structural biology is entering a golden age, where the pace of new structural information is accelerating. We anticipate that future crystallographic and high-resolution cryo-EM studies will define the strategies used by other MAPs to recognize and control the conformation of αβ-tubulin, and may reveal new conformations of αβ-tubulin inside and outside of the microtubule. Reconstitutions of microtubule dynamics are rapidly increasing in complexity and are beginning to reveal how the activities of multiple MAPs can reinforce or antagonize each other (Zanic et al., 2013). More complex reconstitutions are also defining the minimal requirements for creating cellular-scale structures like the mitotic spindle (Bieling et al., 2010; Subramanian et al., 2013). Reconstitutions will also greatly advance the understanding of the dynamics and regulation of microtubule minus ends. As the ever-advancing structural data are integrated with reconstitution data, incorporated into computational models, and correlated with cell biology experiments, a robust, multiscale understanding of microtubule biology will come within reach.     

Concerted actions of DnaA complexes with DNA-unwinding sequences within and flanking replication origin oriC promote DnaB helicase loading

Unwinding of the replication origin and loading of DNA helicases underlie the initiation of chromosomal replication. In Escherichia coli, the minimal origin oriC contains a duplex unwinding element (DUE) region and three (Left, Middle, and Right) regions that bind the initiator protein DnaA. The Left/Right regions bear a set of DnaA-binding sequences, constituting the Left/Right-DnaA subcomplexes, while the Middle region has a single DnaA-binding site, which stimulates formation of the Left/Right-DnaA subcomplexes. In addition, a DUE-flanking AT-cluster element (TATTAAAAAGAA) is located just outside of the minimal oriC region. The Left-DnaA subcomplex promotes unwinding of the flanking DUE exposing TT[A/G]T(T) sequences that then bind to the Left-DnaA subcomplex, stabilizing the unwound state required for DnaB helicase loading. However, the role of the Right-DnaA subcomplex is largely unclear. Here, we show that DUE unwinding by both the Left/Right-DnaA subcomplexes, but not the Left-DnaA subcomplex only, was stimulated by a DUE-terminal subregion flanking the AT-cluster. Consistently, we found the Right-DnaA subcomplex–bound single-stranded DUE and AT-cluster regions. In addition, the Left/Right-DnaA subcomplexes bound DnaB helicase independently. For only the Left-DnaA subcomplex, we show the AT-cluster was crucial for DnaB loading. The role of unwound DNA binding of the Right-DnaA subcomplex was further supported by in vivo data. Taken together, we propose a model in which the Right-DnaA subcomplex dynamically interacts with the unwound DUE, assisting in DUE unwinding and efficient loading of DnaB helicases, while in the absence of the Right-DnaA subcomplex, the AT-cluster assists in those processes, supporting robustness of replication initiation.

The initiation of bacterial DNA replication requires local duplex unwinding of the chromosomal replication origin oriC, which is regulated by highly ordered initiation complexes. In Escherichia coli, the initiation complex contains oriC, the ATP-bound form of the DnaA initiator protein (ATP–DnaA), and the DNA-bending protein IHF (Fig. 1, A and B), which promotes local unwinding of oriC (1, 2, 3, 4). Upon this oriC unwinding, two hexamers of DnaB helicases are bidirectionally loaded onto the resultant single-stranded (ss) region with the help of the DnaC helicase loader (Fig. 1B), leading to bidirectional chromosomal replication (5, 6, 7, 8). However, the fundamental mechanism underlying oriC-dependent bidirectional DnaB loading remains elusive.Open in a separate windowFigure 1Schematic structures of oriC, DnaA, and the initiation complexes. A, the overall structure of oriC. The minimal oriC region and the AT-cluster region are indicated. The sequence of the AT-cluster−DUE (duplex-unwinding element) region is also shown below. The DUE region (DUE; pale orange bars) contains three 13-mer repeats: L-DUE, M-DUE, and R-DUE. DnaA-binding motifs in M/R-DUE, TT(A/G)T(T), are indicated by red characters. The AT-cluster region (AT cluster; brown bars) is flanked by DUE outside of the minimal oriC. The DnaA-oligomerization region (DOR) consists of three subregions called Left-, Middle-, and Right-DOR. B, model for replication initiation. DnaA is shown as light brown (for domain I–III) and darkbrown (for domain IV) polygons (right panel). ATP–DnaA forms head-to-tail oligomers on the Left- and Right-DORs (left panel). The Middle-DOR (R2 box)-bound DnaA interacts with DnaA bound to the Left/Right-DORs using domain I, but not domain III, stimulating DnaA assembly. IHF, shown as purple hexagons, bends DNA >160° and supports DUE unwinding by the DnaA complexes. M/R-DUE regions are efficiently unwound. Unwound DUE is recruited to the Left-DnaA subcomplex and mainly binds to R1/R5M-bound DnaA molecules. The sites of ssDUE-binding B/H-motifs V211 and R245 of R1/R5M-bound DnaA molecules are indicated (pink). Two DnaB homohexamer helicases (light green) are recruited and loaded onto the ssDUE regions with the help of the DnaC helicase loader (cyan). ss, single stranded.The minimal oriC region consists of the duplex unwinding element (DUE) and the DnaA oligomerization region (DOR), which contains specific arrays of 9-mer DnaA-binding sites (DnaA boxes) with the consensus sequence TTA[T/A]NCACA (Fig. 1A) (3, 4). The DUE underlies the local unwinding and contains 13-mer AT-rich sequence repeats named L-, M-, and R-DUE (9). The M/R-DUE region includes TT[A/G]T(A) sequences with specific affinity for DnaA (10). In addition, a DUE-flanking AT-cluster (TATTAAAAAGAA) region resides just outside of the minimal oriC (Fig. 1A) (11). The DOR is divided into three subregions, the Left-, Middle-, and Right-DORs, where DnaA forms structurally distinct subcomplexes (Fig. 1A) (8, 12, 13, 14, 15, 16, 17). The Left-DOR contains high-affinity DnaA box R1, low-affinity boxes R5M, τ1−2, and I1-2, and an IHF-binding region (17, 18, 19, 20). The τ1 and IHF-binding regions partly overlap (17).In the presence of IHF, ATP–DnaA molecules cooperatively bind to R1, R5M, τ2, and I1-2 boxes in the Left-DOR, generating the Left-DnaA subcomplex (Fig. 1B) (8, 17). Along with IHF causing sharp DNA bending, the Left-DnaA subcomplex plays a leading role in DUE unwinding and subsequent DnaB loading. The Middle-DOR contains moderate-affinity DnaA box R2. Binding of DnaA to this box stimulates DnaA assembly in the Left- and Right-DORs using interaction by DnaA N-terminal domain (Fig. 1B; also see below) (8, 12, 14, 16, 21). The Right-DOR contains five boxes (C3-R4 boxes) and cooperative binding of ATP–DnaA molecules to these generates the Right-DnaA subcomplex (Fig. 1B) (12, 18). This subcomplex is not essential for DUE unwinding and plays a supportive role in DnaB loading (8, 15, 17). The Left-DnaA subcomplex interacts with DnaB helicase, and the Right-DnaA subcomplex has been suggested to play a similar role (Fig. 1B) (8, 13, 16).In the presence of ATP–DnaA, M- and R-DUE adjacent to the Left-DOR are predominant sites for in vitro DUE unwinding: unwinding of L-DUE is less efficient than unwinding of the other two (Fig. 1B) (9, 22, 23). Deletion of L-DUE or the whole DUE inhibits replication of oriC in vitro moderately or completely, respectively (23). A chromosomal oriC Δ(AT-cluster−L-DUE) mutant with an intact DOR, as well as deletion of Right-DOR, exhibits limited inhibition of replication initiation, whereas the synthetic mutant combining the two deletions exhibits severe inhibition of cell growth (24). These studies suggest that AT-cluster−L-DUE regions stimulate replication initiation in a manner concerted with Right-DOR, although the underlying mechanisms remain elusive.DnaA consists of four functional domains (Fig. 1B) (4, 25). Domain I supports weak domain I–domain I interaction and serves as a hub for interaction with various proteins such as DnaB helicase and DiaA, which stimulates ATP–DnaA assembly at oriC (26, 27, 28, 29, 30). Two or three domain I molecules of the oriC–DnaA subcomplex bind a single DnaB hexamer, forming a stable higher-order complex (7). Domain II is a flexible linker (28, 31). Domain III contains AAA+ (ATPase associated with various cellular activities) motifs essential for ATP/ADP binding, ATP hydrolysis, and DnaA–DnaA interactions in addition to specific sites for ssDUE binding and a second, weak interaction with DnaB helicase (1, 4, 8, 10, 19, 25, 32, 33, 34, 35). Domain IV bears a helix-turn-helix motif with specific affinity for the DnaA box (36).As in typical AAA+ proteins, a head-to-tail interaction underlies formation of ATP–DnaA pentamers on the DOR, where the AAA+ arginine-finger motif Arg285 recognizes ATP bound to the adjacent DnaA protomer, promoting cooperative ATP–DnaA binding (Fig. 1B) (19, 32). DnaA ssDUE-binding H/B-motifs (Val211 and Arg245) in domain III sustain stable unwinding by directly binding to the T-rich (upper) strand sequences TT[A/G]T(A) within the unwound M/R-DUE (Fig. 1B) (8, 10). Val211 residue is included in the initiator-specific motif of the AAA+ protein family (10). For DUE unwinding, ssDUE is recruited to the Left-DnaA subcomplex via DNA bending by IHF and directly interacts with H/B-motifs of DnaA assembled on Left-DOR, resulting in stable DUE unwinding competent for DnaB helicase loading; in particular, DnaA protomers bound to R1 and R5M boxes play a crucial role in the interaction with M/R-ssDUE (Fig. 1B) (8, 10, 17). Collectively, these mechanisms are termed ssDUE recruitment (4, 17, 37).Two DnaB helicases are thought to be loaded onto the upper and lower strands of the region including the AT-cluster and DUE, with the aid of interactions with DnaC and DnaA (Fig. 1B) (25, 38, 39). DnaC binding modulates the closed ring structure of DnaB hexamer into an open spiral form for entry of ssDNA (40, 41, 42, 43). Upon ssDUE loading of DnaB, DnaC is released from DnaB in a manner stimulated by interactions with ssDNA and DnaG primase (44, 45). Also, the Left- and Right-DnaA subcomplexes, which are oriented opposite to each other, could regulate bidirectional loading of DnaB helicases onto the ssDUE (Fig. 1B) (7, 8, 35). Similarly, recent works suggest that the origin complex structure is bidirectionally organized in both archaea and eukaryotes (1, 46). In Saccharomyces cerevisiae, two origin recognition complexes containing AAA+ proteins bind to the replication origin region in opposite orientations; this, in turn, results in efficient loading of two replicative helicases, leading to head-to-head interactions in vitro (46). Consistent with this, origin recognition complex dimerization occurs in the origin region during the late M-G1 phase (47). The fundamental mechanism of bidirectional origin complexes might be widely conserved among species.In this study, we analyzed various mutants of oriC and DnaA in reconstituted systems to reveal the regulatory mechanisms underlying DUE unwinding and DnaB loading. The Right-DnaA subcomplex assisted in the unwinding of oriC, dependent upon an interaction with L-DUE, which is important for efficient loading of DnaB helicases. The AT-cluster region adjacent to the DUE promoted loading of DnaB helicase in the absence of the Right-DnaA subcomplex. Consistently, the ssDNA-binding activity of the Right-DnaA subcomplex sustained timely initiation of growing cells. These results indicate that DUE unwinding and efficient loading of DnaB helicases are sustained by concerted actions of the Left- and Right-DnaA subcomplexes. In addition, loading of DnaB helicases are sustained by multiple mechanisms that ensure robust replication initiation, although the complete mechanisms are required for precise timing of initiation during the cell cycle.     

Mechanisms of spindle positioning

Accurate positioning of spindles is essential for asymmetric mitotic and meiotic cell divisions that are crucial for animal development and oocyte maturation, respectively. The predominant model for spindle positioning, termed “cortical pulling,” involves attachment of the microtubule-based motor cytoplasmic dynein to the cortex, where it exerts a pulling force on microtubules that extend from the spindle poles to the cell cortex, thereby displacing the spindle. Recent studies have addressed important details of the cortical pulling mechanism and have revealed alternative mechanisms that may be used when microtubules do not extend from the spindle to the cortex.Mitotic and meiotic spindles are precisely positioned within eukaryotic cells for several reasons. In animal cells, spindle position determines the location of contractile ring assembly (Green et al., 2012). Thus, placing a spindle in the center of the cell will result in daughter cells of equal size, whereas positioning the spindle asymmetrically results in daughter cells of different sizes. In oocytes, the extreme asymmetrical positioning of the meiotic spindle allows expulsion of three fourths of the chromosomes into two tiny polar bodies while preserving most of the cytoplasm in the egg for the developing zygote. In polarized cells, where proteins and RNAs are asymmetrically distributed before division, the orientation of the spindle relative to the polarity axis determines whether the daughter cells will have the same or different developmental fates. An excellent review of the developmental context of spindle positioning is provided by Morin and Bellaïche (2011). In budding yeast (Slaughter et al., 2009) and plants (Rasmussen et al., 2011), the site of cytokinesis is determined before the spindle forms. In these organisms, the spindle must be oriented relative to the predetermined division plane to ensure that both daughter cells receive a complete chromosome complement.The majority of research on the mechanisms of spindle positioning has focused on cell types that have “astral” microtubules. Astral microtubules have minus ends embedded in the spindle poles and plus ends extending outward, away from the spindle toward the cell cortex (Fig. 1, A and B). Astral microtubules have been proposed to mediate spindle positioning by generating pulling forces at the cortex or pulling forces against the cytoplasm. The minus ends of astral microtubules are embedded in the pericentriolar material that surrounds the centrioles of animal cells or in the spindle pole bodies of fungi. This attachment is essential for pulling forces on the astral microtubules to move the spindle. However, late stage oocytes of several animal phyla and all cells of flowering plants lack centrioles and lack obvious astral microtubules. Thus, these cell types have evolved alternative spindle positioning mechanisms. Here we review recent advances in both astral and nonastral spindle positioning mechanisms.Open in a separate windowFigure 1.Mitotic spindle movements in the C. elegans zygote. (A) Schematic diagram of a single-celled C. elegans embryo showing cortical pulling by cytoplasmic dynein during pronuclear centration and rotation. The nuclei move toward the anterior (left) so that the spindle assembles in the center of the embryo. (B) Schematic diagram of a single-celled C. elegans embryo showing cortical pulling by dynein during anaphase. The spindle moves to the posterior (right) so that cytokinesis generates two cells of different sizes. The squares highlight a dynein molecule that pulls toward the posterior before spindle displacement, then pulls toward the anterior after spindle displacement. (C) Schematic drawing of cytoplasmic pulling that contributes to centering the pronuclei. (D) Illustration of a spindle that was centered at metaphase but in which both poles moved all the way to the posterior end of the embryo. This occurs in zyg-8 mutants (Gönczy et al., 2001), cls-1,2 (RNAi) embryos (Espiritu et al., 2012), and zyg-9(ts) mutants shifted to a nonpermissive temperature at metaphase (Bellanger et al., 2007), possibly because astral microtubules are too short to reach force generators that would pull toward the anterior.

Cortical versus cytoplasmic pulling

The most prominent model of spindle positioning involves a cortical pulling mechanism. In this model, the minus end–directed microtubule motor protein, cytoplasmic dynein, is attached to the cell cortex and exerts pulling forces on the plus ends of astral microtubules that reach the cortex. In the single-celled Caenorhabditis elegans embryo at early prophase, complexes of GPR-1,2 (G protein regulator) and LIN-5 (abnormal cell lineage), positive regulators of cytoplasmic dynein, are more concentrated at the anterior cortex of the embryo (Fig. 1 A; Park and Rose, 2008), resulting in greater net pulling force toward the anterior. This results in net movement of the pronuclei to the center of the embryo and rotation of the centrosome–pronuclear complex so that the metaphase spindle forms in the center of the embryo with its poles oriented along the anterior-posterior axis of the embryo (Fig. 1 B). During metaphase and early anaphase, GPR-1,2 and LIN-5 become more concentrated at the posterior end of the embryo, resulting in movement of the spindle toward the posterior so that cytokinesis generates daughter cells of different sizes (Fig. 1 B; Grill et al., 2001, 2003). Depending on the relative distribution of active force generators at the cortex, this mechanism can also lead to centering the spindle within the cell to allow symmetric cytokinesis as occurs in HeLa cells (Kiyomitsu and Cheeseman, 2012) and LLC-Pk1 cells (Collins et al., 2012). Cortical pulling might involve pulling on the sides of microtubules that bend as they approach the cortex (Fig. 1 A, 1) or end-on interactions that require coupling of microtubule depolymerization with pulling (Fig. 1 A, 2), as occurs at kinetochores during anaphase A (McIntosh et al., 2010).Cortical pulling differs from a cytoplasmic pulling mechanism most clearly proposed by Kimura and Kimura (2011) and diagrammed in Fig. 1 C. In this cytoplasmic pulling mechanism, the viscous drag on membranous organelles transported toward the minus ends of astral microtubules by cytoplasmic dynein generates a force in the opposite direction, toward the cortex. Depletion of RAB-5, RAB-7, or RILP-1 (RAB-7 interacting lysosomal protein homologue) blocks dynein-dependent organelle transport and slows the velocity of pronuclear centration in C. elegans without affecting other dynein-dependent movements (Kimura and Kimura, 2011). Elimination of cortical pulling by depleting GPR-1,2 also slows pronuclear centration (Park and Rose, 2008), which suggests that cortical pulling and cytoplasmic pulling each contribute 50% of the velocity of pronuclear centration. Unlike end-on cortical pulling, cytoplasmic pulling force is proportional to the length of the astral microtubules because more organelles will be transported on a long microtubule than a short microtubule (Fig. 1 C). This generates a self-centering mechanism as the length of the astral microtubules equalize when the pronuclei reach the center of the zygote (Fig. 1 A; Hamaguchi and Hiramoto, 1986). Cytoplasmic pulling may predominate in very large zygotes where astral microtubules clearly do not reach the cortex but where both pronuclei and the mitotic spindle are centered (Mitchison et al., 2012).

Evidence for cortical pulling.

The cortical pulling mechanism requires that microtubules extend from the spindle to the cortex and form a contiguous structure that is mechanically robust enough that the force generators do not pull the minus ends of the microtubules out of the spindle pole or cause the plasma membrane to buckle inward. Experimental evidence for this contiguous mechanical linkage comes from experiments in oocytes of the marine annelid, Chaetopterus. Insertion of a glass needle into the meiotic spindle allowed pulling the spindle away from the cortex, which caused inward buckling of the cortex. Further pulling resulted in sudden release of the spindle from the cortex, restoration of cortical shape, and concomitant disappearance of a birefringent aster extending between the spindle and cortex (Lutz et al., 1988). The second requirement for a cortical pulling mechanism is that force is generated at the cortex. Using a laser to cut the central spindle of an early anaphase C. elegans embryo, Grill et al. (2001) showed that spindle poles are pulled from outside the spindle rather than pushed from inside the spindle during posterior spindle displacement. Fragmentation of centrosomes with a laser (Grill et al., 2003) revealed that astral microtubules freed from the spindle move outward toward the cortex. Either cortical pulling or cytoplasmic pulling could explain this result; however, the asymmetric distribution of fragment velocities is controlled by proteins that are localized at the cortex, GPR-1,2 (Grill et al., 2003) and LET-99 (lethal-99; Krueger et al., 2010). In a key experiment, Redemann et al. (2010) showed that microtubule plus ends pull tubular invaginations of the plasma membrane inward when cortical stiffness is partially reduced. This experiment showed that astral microtubules are pulling on the cortex during spindle displacement, which would not occur if forces were generated by movement of cytoplasmic organelles along the sides of microtubules.

End-on versus side-on microtubule–cortex interactions.

How do microtubules interact with force generators at the cortex? Astral microtubules might polymerize to the cortex then bend along it so that cortical motors interact with the side of the microtubule to generate force (Fig. 1 A, 1). This type of interaction would be consistent with the in vitro gliding motility of microtubules generated by cytoplasmic dynein immobilized on glass coverslips (Paschal et al., 1987; Vallee et al., 1988) and is the only type of dynein-dependent cortical microtubule interaction observed in budding yeast (Adames and Cooper, 2000). Studies of C. elegans embryos, however, support end-on microtubule contacts (Fig. 1 A, 2) as the functional contact with cortical force generators for posterior displacement of the early anaphase spindle. Live imaging of YFP-tubulin in optical sections at the surface of the embryo during anaphase revealed dots rather than lines, indicating that the microtubules contacting the cortex are <200 nm in length (Labbé et al., 2003; Kozlowski et al., 2007). When short microtubule fragments are generated by ectopic katanin activity, dynein-dependent gliding of microtubule “lines” on the cortex is frequently observed (Gusnowski and Srayko, 2011), indicating that cortical dynein is capable of moving microtubules along the cortex via side-on interactions in wild-type embryos but that it does not during posterior displacement of the anaphase spindle. A likely explanation comes from the finding that astral microtubule plus ends undergo catastrophe (switch to depolymerization) on average 1.4 s after polymerizing to the cortex (Kozlowski et al., 2007). Thus astral microtubule plus ends do not have time to polymerize along the cortex to establish extensive side-on contacts. Support for this idea comes from depletion of the conserved plasma membrane protein EFA-6 (exchange factor for Arf) from the C. elegans embryo. In the absence of EFA-6, the residence time of microtubule plus ends at the cortex increases fivefold, astral microtubules form extensive lateral contacts with the cortex, and centrosomes exhibit movements consistent with excessive dynein-dependent cortical pulling (O’Rourke et al., 2010). Side-on contacts of astral microtubules with the cortex occur later during telophase in the wild-type C. elegans embryo (Kozlowski et al., 2007), but the nature of this switch has not been addressed.The two distinct activities of cytoplasmic dynein, end-on pulling and walking along the side of a microtubule, have been genetically separated in C. elegans. Cortical pulling forces during early anaphase require the redundant cortical dynein activators GPR-1 and -2 (Grill et al., 2003), which bind to LIN-5 (Gotta et al., 2003). GPR-1,2/LIN-5 is anchored in the plasma membrane via the myristoyl and palmitoyl lipid modifications of the redundant Gα proteins GOA-1 and GPA-16 (Gotta et al., 2003; Park and Rose, 2008; Kotak et al., 2012). The complex of GPR-1 and LIN-5 interacts with the dynein light chain DYRB-1 (Couwenbergs et al., 2007) and the dynein regulator LIS-1 (human lissencephaly gene related; Nguyen-Ngoc et al., 2007). GPR-1 and -2, however, are not required for dynein-dependent gliding of severed microtubule fragments along the cortex (Gusnowski and Srayko, 2011), dynein-dependent transport of membranous organelles along the sides of microtubules (Kimura and Kimura, 2011), dynein-dependent positioning of the acentriolar C. elegans meiotic spindle (van der Voet et al., 2009), or dynein-dependent centration of the male pronucleus (Kimura and Kimura, 2011). These GPR-independent activities of cytoplasmic dynein likely do not require end-on pulling.Recently, end-on pulling by cytoplasmic dynein has been reconstituted in vitro with a purified preparation of artificially dimerized budding yeast cytoplasmic dynein. Laan et al. (2012) immobilized purified cytoplasmic dynein on microfabricated barriers and observed the interaction of centrosome-nucleated microtubules as they approached these dynein-coated barriers. Microtubule plus ends hitting a dynein-coated barrier switched to catastrophe with high frequency but the microtubule depolymerization rate after the catastrophe was reduced. The result was an extended period of interaction between the depolymerizing plus end and the dynein-coated barrier. Plus-end depolymerization pulled the centrosome toward the barrier, and in similar reactions the pulling force was measured as high as 5 pN. Side-on interactions with the barrier were not observed. Whereas ATP was required for this end-on pulling, it is not clear if the energy source is ATP hydrolysis–driven stepping by dynein or if the energy source is GTP hydrolysis–driven depolymerization of the microtubule. In the latter case, ATP might only be required to prevent rigor binding of dynein to the microtubule. Indeed, an artificial rigor binding of a streptavidin-coated bead to a depolymerizing biotinylated microtubule plus end resulted in a pulling force that is restricted to an extremely short distance (Grishchuk et al., 2005). In vitro reconstitution of pulling force coupled to depolymerizing microtubule plus ends was first demonstrated with beads coated with kinesin-1 or a nonmotile kinesin chimera (NK350; Lombillo et al., 1995). Like barrier-bound cytoplasmic dynein, kinesin-coated beads slowed the depolymerization rate of microtubule plus ends, whereas NK350-coated beads enhanced the depolymerization rate of bound plus ends. ATP enhanced depolymerization-coupled pulling for both kinesin-1 and NK350, just as it did for barrier-bound cytoplasmic dynein. The in vitro pulling reaction reconstituted by Laan et al. (2012) seems unlikely to be the same reaction that pulls on plus ends in the anaphase budding yeast cell, as these are through side-on interactions (Adames and Cooper, 2000). In vitro reconstitution of a GPR/LIN-5–dependent, end-on pulling reaction with purified metazoan cytoplasmic dynein may reveal mechanisms acting on spindles in vivo.Another interesting contrast between end-on versus side-on cortical pulling reactions is suggested by differences in the dependence on cortical F-actin. F-actin is required for cortical rigidity to prevent end-on microtubule contacts from pulling membrane tubules inward instead of moving the spindle pole outward (Redemann et al., 2010). Side-on pulling by cortical dynein in budding yeast, however, does not require F-actin (Theesfeld et al., 1999; Heil-Chapdelaine et al., 2000a). The curvature of microtubules gliding on the bud cortex indicates that the microtubule is engaged with multiple dynein molecules distributed over several microns of cortex, and this distribution of force might allow effective pulling against a less rigid cortex. Alternatively, rigidity of the yeast plasma membrane might be mediated by oligomers of BAR domain proteins like Num1 (nuclear migration; Tang et al., 2012) or eisosomes (Walther et al., 2006; Olivera-Couto et al., 2011), by osmotic pressure, or by attachment of the plasma membrane to the cell wall.

Why the spindle is not pulled all the way to the cortex with more active force generators.

What prevents the C. elegans centrosome–pronuclear complex from moving all the way to the anterior cortex where the concentration of cortical force generators is highest during prophase (Fig. 1 A), and what prevents the spindle from moving all the way to the posterior cortex, which has the highest concentration of active force generators during anaphase (Fig. 1, B and D)? Increasing the concentration of GPR-1,2/LIN-5 at the anterior cortex causes pronuclei to move further toward the anterior, but they still do not crash into the anterior cortex (Panbianco et al., 2008). During metaphase/anaphase (Fig. 1 B), weak cortical pulling on the anterior aster might oppose strong pulling on the posterior aster. Supporting this idea, spindle severing results in the posterior aster moving further posterior than when the spindle is intact (Grill et al., 2001), but the posterior pole still does not move all the way to the posterior cortex. Monopolar spindles move toward the posterior in a GPR1,2-dependent manner but then reverse direction and oscillate along the anterior-posterior axis (Krueger et al., 2010). Laan et al. (2012) found that centrosome-nucleated microtubule asters could accurately self-center within microchambers whose walls are coated with dynein. Their mathematical modeling suggested that self-centering was achieved because of a balance between cortical pulling by dynein and pushing by polymerizing microtubules (Dogterom and Yurke, 1997) that have not yet engaged a dynein molecule. Thus, cortical pushing by astral microtubules might buffer cortical pulling in vivo. Grill and Hyman (2005) suggested another simple solution. When a spindle pole moves to the posterior, it passes a subset of cortical force generators that were initially pulling toward the posterior (Fig. 1 B, box). After passage of the spindle pole, these force generators pull toward the anterior. Failure in any of these buffering mechanisms might explain why both spindle poles move to the posterior cortex (Fig. 1 D) in mutants that have either short astral microtubules or fewer astral microtubules reaching the cortex (Gönczy et al., 2001; Bellanger et al., 2007; Espiritu et al., 2012).Recent experiments in HeLa and LLC-Pk1 cells have revealed a more sophisticated feedback mechanism that effectively centers the mitotic spindle. Kiyomitsu and Cheeseman (2012) found that HeLa cell mitotic spindles oscillate back and forth along their pole-to-pole axis. They found that dynein/dynactin formed a lateral crescent on the cortex when the spindle was far from that lateral cortex (Fig. 2 A). As the spindle moved toward the dynein crescent, the dynein crescent disappeared as the spindle approached and a new crescent appeared on the opposite lateral cortex (Fig. 2 C). They found that polo kinase 1 (Plk1), which is concentrated on spindle poles, causes dynein/dynactin to dissociate from the LGN–NuMA–Gαi complex (Leu-Gly-Asn repeat enriched protein–nuclear mitotic apparatus protein–Gα; homologues of GPR-1,2–LIN-5–Gα), which explains why the dynein crescent disappears once the spindle pole gets close to the cortex. They also found that the GTP-Ran gradient produced by chromosome-bound RCC1 (regulator of chromosome condensation) was responsible for inhibiting LGN–NuMA association with the cortex, and hence dynein, above the central spindle (Fig. 2). The RCC1 pathway explains why the spindles move only along their pole–pole axis. The two pathways combine to center the spindle in two different axes. Similar spindle oscillations with dynein/dynactin crescents forming only when the spindle pole is far from the cortex were reported in LLC-Pk1 epithelial cells (Collins et al., 2012). Because spindles are small relative to the two-dimensional flattened area of these cells, the role of this pathway in centering spindles to allow symmetric cytokinesis is much more obvious than in HeLa cells.Open in a separate windowFigure 2.How to center a spindle. Schematic diagram of a metaphase HeLa cell where the spindle oscillates along its pole–pole axis to maintain a centered position to allow symmetric cytokinesis. (A) When the left spindle pole is close to the cortex, Plk1 on the pole (red) causes dynein (green) to dissociate from LGN–NuMA complexes (purple; human homologues of GPR-1,2/LIN-5). (B and C) The spindle moves to the right because of the higher concentration of LGN–NuMA–dynein complexes on the right cortex. When chromosomes are close to the cortex as in A, the GTP-Ran gradient from the chromosomes causes dissociation of LGN/NuMA from the cortex. This second system centers the spindle in the axis perpendicular to the pole–pole axis.There are normal situations where movement of one spindle pole all the way to the cortex occurs. This exaggerated movement that results in one set of astral microtubules being splayed onto the cortex is observed in fourth cleavage sea urchin embryos (Holy and Schatten, 1991) and for the female meiotic spindles of Chaetopterus (Lutz et al., 1988) and Spisula solidissima (Pielak et al., 2004).

The budding yeast model.

The S. cerevisiae mitotic spindle is positioned relative to a preformed bud neck by two sequential and partially redundant cortical pulling pathways to ensure that chromosomes are deposited into both mother and daughter cells. Spindle positioning is also monitored by a budding yeast–specific checkpoint (Caydasi et al., 2010). Deletion of genes in any one positioning pathway results in a viable yeast strain, whereas double mutants bearing deletions of genes in both positioning pathways or of genes in one positioning pathway plus the checkpoint results in lethality (Miller et al., 1998). In the early pathway, the microtubule plus-end tip tracking protein, Bim1 (binding to microtubules; EB1 homologue), binds to the yeast-specific adaptor protein Kar9 (karyogamy; Korinek et al., 2000; Lee et al., 2000; Miller et al., 2000), which binds to the yeast myosin V (Myo2; Yin et al., 2000). Myosin V transports the growing microtubule plus end toward the bud tip on polarized actin cables (Hwang et al., 2003). This results in a unique “sweeping” or “pivoting” of the growing astral microtubule toward the bud neck (Fig. 3 A; Adames and Cooper, 2000). Because Bim1 only binds to growing microtubule plus ends (Zimniak et al., 2009), this mechanism alone cannot bring the spindle to the bud neck because the polymerizing astral microtubule would push the spindle back into the mother cell. To prevent the astral microtubules from becoming too long, the kinesin-8 family member, Kip3, passively tracks the plus end until the plus end reaches the bud cortex (Fig. 3 A). Kip3 then switches to a plus-end depolymerase and shortens the astral microtubule to pull the spindle to the bud neck (Fig. 3, B and C; Gupta et al., 2006). Purified Kip3 has unique biochemical properties that contribute to its in vivo function. In vitro, Kip3 accumulates at plus-end tips via its plus end–directed motor activity. Longer microtubules accumulated more Kip3 at their plus ends than short microtubules simply because there are more lateral binding sites on a long microtubule and Kip3 switches to a plus-end depolymerase only when the microtubule has reached a threshold length (Varga et al., 2009). Plus-end pulling by the fission yeast homologue of Kip3 has also been reconstituted in vitro (Grissom et al., 2009). More recent work suggests that Kip3-dependent cortical pulling requires the concerted action of the cortical protein, Bud6 (BUD site selection), and the plus end tracker, Bim1, as well as cytoplasmic dynein (ten Hoopen et al., 2012).Open in a separate windowFigure 3.Spindle positioning in budding yeast. Schematic diagram of the two sequential spindle positioning pathways of budding yeast. In the early pathway (A–C), myosin V transports the plus end of an astral microtubule toward the bud tip on a polarized actin cable. Once the plus end has reached the bud cortex, the plus-end depolymerase, KIP3, is activated to allow pulling of the spindle pole toward the bud neck. In the late pathway (D–F), the plus end–directed microtubule motor Kip2 transports dynein to the plus ends of microtubules via the adaptor protein Bik1 (D). Dynein can be targeted to plus ends by two additional Bik1-dependent mechanisms (see text). When dynein reaches the bud cortex on a polymerizing microtubule plus end (E), contact with the cortical protein Num1 allows dynein to pull the spindle toward the bud (F). During early anaphase (D and E), dynein is not loaded onto microtubules in the mother cell. During late anaphase (F), dynein is loaded on microtubules in the mother cell to prevent movement of the spindle all the way into the bud.The late pathway is initiated when the yeast-specific dynein inhibitor She1 (sensitivity to high expression) is removed from astral microtubules at the metaphase–anaphase transition (Woodruff et al., 2009). She1 appears to act specifically by preventing recruitment of dynactin to microtubules (Bergman et al., 2012) and by inhibiting dynein motility (Markus et al., 2012). In the late pathway, cytoplasmic dynein is targeted to growing microtubule plus ends by the plus-end tracking protein Bik1 (bilateral karyogamy defect; Sheeman et al., 2003), which itself is targeted to plus ends by three partially redundant mechanisms. In the first mechanism, Bik1 is transported as cargo by the plus end–directed kinesin Kip2 (Carvalho et al., 2004). Cytoplasmic dynein is thus transported as cargo to the bud cortex by Bik1–Kip2 complexes (Fig. 3 D). In the absence of Kip2, Bik1 (and therefore dynein) can still track growing microtubule plus ends either through a second mechanism that requires the C-terminal tyrosine residue of α-tubulin or a third mechanism that requires the plus end–tracking protein Bim1 (Caudron et al., 2008). This is partially consistent with results of reconstitution experiments with purified proteins showing that the Bik1 homologue, CLIP170, tracks growing plus ends through a mechanism that involves binding to both the Bim1 homologue, EB1, and the C-terminal tyrosine-containing motif of α-tubulin (Bieling et al., 2008). When a microtubule plus end carrying dynein contacts the yeast-specific cortical protein Num1, Num1 apparently stimulates off-loading of the dynein tail onto the cortex so that the dynein motor domains engage the microtubule in a cortical pulling reaction (Fig. 3, E and F). Deletion of Num1 results in accumulation of inactive dynein at plus ends (Lee et al., 2003; Sheeman et al., 2003; Markus and Lee, 2011). In contrast with the GPR-dependent end-on cortical interactions observed in C. elegans, dynein-dependent cortical pulling is mediated by lateral sliding of astral microtubules along the yeast bud cortex (Fig. 3, E and F; Adames and Cooper, 2000). Also, unlike cortical GPR/LIN-5 in animal cells, cortical Num1 is distributed in patches throughout both mother and daughter cells (Heil-Chapdelaine et al., 2000b) and even participates in mitochondrial positioning and fission throughout the cell (Cerveny et al., 2007; Hammermeister et al., 2010). If there is no asymmetrically distributed cortical activator of dynein, how is dynein-mediated pulling directed specifically toward the bud cortex?

Why both spindle poles are not pulled to the same cortex in budding yeast.

The budding yeast spindle, rather than the cortex, is asymmetrical. At metaphase, Kar9 is asymmetrically localized on the spindle pole oriented toward the bud and on the plus ends of astral microtubules emanating from that bud-proximal spindle pole (Liakopoulos et al., 2003). This alone would explain why only one spindle pole moves toward the bud but then leaves the question of how Kar9 asymmetry is established. Cepeda-García et al. (2010) found that Kar9 at spindle poles became symmetrical and reduced in concentration after depolymerization of actin cables, depolymerization of microtubules, or disruption of the myosin V–Kar9 interaction. When microtubules were repolymerized, Kar9 was initially symmetrical on both spindle poles and quickly repolarized onto the first microtubule to make a functional cortical contact. These results suggested a positive feedback loop in which functional cortical pulling by Bim1–Kar9–Myo2 complexes causes loading of additional Kar9. Cytoplasmic dynein also accumulates preferentially on the plus-end tips that reach the bud neck first (Fig. 3, D and E) and on the bud-proximal spindle pole during metaphase. The asymmetrical accumulation of dynein on the bud-proximal microtubules and bud-proximal spindle pole requires kinases that are found at the bud neck. Thus, the asymmetry of dynein localization may be generated by a positive feedback loop, as suggested for Kar9 asymmetry. After the spindle is pulled into the bud neck, during anaphase, dynein becomes symmetrical on the plus ends emanating from both poles (Fig. 3 F; Grava et al., 2006). This regulation prevents both poles from moving toward the bud during metaphase and then prevents the spindle from being pulled all the way into the bud during anaphase. Asymmetric localization of dynein on spindle poles or microtubule plus ends has not yet been reported in animal cells.

Other spindle positioning mechanisms

In the examples of the C. elegans and budding yeast mitotic spindles, long astral microtubules are in contact with the cell cortex. In cells where spindles have no astral microtubules, other mechanisms must be at work. Female meiotic spindles are universally positioned with one spindle pole contacting the oocyte cortex so that one set of chromosomes can be eliminated in a polar body through an extremely asymmetrical division (Fabritius et al., 2011). Female meiotic spindles of at least three animal phyla (Chordata, Nematoda, and Arthropoda), however, have no centrioles in their spindle poles and no apparent astral microtubules. Work in C. elegans and mice suggests that different species have evolved different mechanisms for acentriolar meiotic spindle positioning.

Parallel metaphase meiotic spindles.

The metaphase I and metaphase II spindles of C. elegans (Fig. 4 A) and the metaphase II spindle of mouse (Fig. 4 B) are positioned at the cortex in a parallel orientation, with both poles equidistant from the cortex. In C. elegans, this parallel cortical positioning requires microtubules, kinesin-1, and a worm-specific kinesin-1–binding partner, KCA-1 (Yang et al., 2003, 2005), but is independent of F-actin (Yang et al., 2003). Kinesin-1 and microtubules are also required to move the nucleus to the cortex before germinal vesicle breakdown and to drive transport of yolk granules inward from the cortex, which results in a circular streaming pattern. It has been suggested that kinesin-1 may only move the germinal vesicle to the cortex and that an additional, unidentified pathway moves the spindle over the remaining distance to the cortex and establishes the parallel orientation (McNally et al., 2010). The mouse metaphase II spindle is maintained in a similar parallel orientation at the cortex, but this positioning requires the actin nucleator ARP2/3. ARP2/3 also drives streaming of actin filaments and cytoplasm in a pattern that has been proposed to push the spindle into the cortex to maintain parallel cortical position (Fig. 4 B; Yi et al., 2011).Open in a separate windowFigure 4.A plethora of nonastral spindle positioning mechanisms. (A) Metaphase C. elegans meiotic spindles are positioned in a parallel orientation at the cortex by microtubules and kinesin-1. (B) The mouse metaphase II spindle may be positioned by actin-dependent cytoplasmic streaming. Pole-first migration of the mouse meiosis I spindle to the cortex may be mediated by cargo transport on parallel actin filaments by spindle pole–bound myosin II (C), myosin II–based contraction of anti-parallel actin filaments (D), or pushing forces generated by polymerizing actin filaments nucleated by formin molecules on the spindle (E) or nucleated by formin molecules on the cortex (F). Red arrows indicate the pointed ends of actin filaments. (G) One spindle pole of the early anaphase C. elegans meiotic spindle may be transported to the cortex as cargo by dynein on polarized cytoplasmic microtubules.

Pole first migration of the mouse meiosis I spindle.

Unlike the C. elegans germinal vesicle, the mouse germinal vesicle is centered in the egg at germinal vesicle breakdown. Thus, the meiosis I spindle assembles near the center of the egg then migrates in a pole-first orientation to the nearest cortex so that it never adopts a parallel orientation (Fig. 4 C). This migration requires F-actin (Verlhac et al., 2000) and the actin nucleators Formin 2 (Dumont et al., 2007), Spire 1 and Spire 2 (Pfender et al., 2011), and ARP2/3 (Sun et al., 2011), but the mechanism of movement remains unclear. Schuh and Ellenberg (2008) demonstrated the existence of actin bundles extending between the spindle and an invagination of the cortex during spindle migration. The invagination indicated a pulling mechanism, and Schuh and Ellenberg (2008) suggested that myosin II on the spindle poles might walk on a discontinuous actin network with barbed ends oriented toward the cortex (Fig. 4 C). In support of this model, the Rho kinase inhibitor ML7 eliminated spindle pole staining by an antibody specific for phosphorylated myosin regulatory light chain and blocked spindle migration (Schuh and Ellenberg, 2008). However, Li et al. (2008) found that the myosin ATPase inhibitor, blebbistatin, had no effect on spindle migration even though it completely blocked polar body extrusion (cytokinesis). Because myosin II typically acts by forming bipolar thick filaments that exert contractile force on antiparallel actin filaments, a myosin II–based model would make more sense if myosin II was concentrated on antiparallel actin bundles extending between the spindle and cortex (Fig. 4 D). Myosin V is more typically associated with the transport of cargo on uniformly oriented actin filaments, and a recent study has shown that myosin V drives transport of secretory vesicles outward toward the cortex in germinal vesicle stage oocytes (Schuh, 2011). This study at least suggests that the cytoplasmic actin meshwork has a net polarity with barbed ends toward the cortex, a prerequisite for a myosin cargo transport model (Fig. 4 C).Li et al. (2008) proposed a completely different mechanism in which F-actin nucleated near the chromosomes generates a cloud of F-actin that pushes the spindle toward the cortex (Fig. 4 E) in a manner analogous to Listeria monocytogenes motility (Lambrechts et al., 2008). Support for this pushing model comes from imaging of an actin cloud behind the migrating spindle and localization of Formin 2 around the spindle (Li et al., 2008). In addition, Formin 2 overexpression causes invaginations in the nuclear envelope, which is consistent with inward pushing from the cortex, rather than protrusions that would be consistent with pulling forces from the cortex (Azoury et al., 2011). Formin 2 is symmetrical around the cortex during prophase but clears from the cortex in front of the migrating spindle, which is consistent with nucleation-based pushing from behind (Fig. 4 F; Azoury et al., 2011). As previously mentioned, cortical stiffness is a prerequisite for cortical pulling mechanisms because pulling on an unsupported plasma membrane should cause the membrane to invaginate inward instead of the spindle moving outward. Strikingly, cortical stiffness of the mouse oocyte decreases sixfold during spindle migration (Larson et al., 2010). Clearly, more work is required to resolve the mechanism of pole-first spindle migration in the mouse oocyte.

Rotation of the parallel metaphase spindle to a perpendicular anaphase spindle.

Activation of the anaphase-promoting complex results in rotation of the parallel metaphase meiotic spindle to a perpendicular orientation in both meiotic divisions of C. elegans, whereas fertilization induces rotation during meiosis II in mouse. In kinesin-1–depleted C. elegans embryos, the metaphase I spindle is far from the cortex and initiates pole-first migration to the cortex at the same time that wild-type rotation initiates (Yang et al., 2005). Both spindle rotation and late spindle migration in a kinesin mutant require cytoplasmic dynein (Ellefson and McNally, 2009, 2011). These results indicate that spindle rotation is simply migration of one spindle pole toward the cortex (analogous to spindle migration during mouse meiosis I). However, unlike dynein-dependent migration of one spindle pole toward the cortex during C. elegans mitosis or HeLa cell mitosis, C. elegans meiotic spindle rotation does not require GPR-1,2 (van der Voet et al., 2009). One possible model for C. elegans meiotic spindle rotation is that cytoplasmic dynein, which accumulates on both spindle poles just before and during rotation, transports one spindle pole as cargo on cytoplasmic microtubules with minus ends anchored at the cortex (Fig. 4 G). The orientation of these microtubules is inferred from the direction of kinesin-dependent yolk granule movement (McNally et al., 2010) and hook decoration in Xenopus laevis oocytes (Pfeiffer and Gard, 1999). This model is essentially the same as the myosin cargo transport model proposed for mouse meiosis I (Fig. 4 C). Rotation of the mouse meiosis II spindle is actin dependent (Maro et al., 1984) and myosin II dependent (Matson et al., 2006; Wang et al., 2011), but the mechanism is unknown.

Unifying themes and future directions

Work in HeLa cells and budding yeast suggests that negative feedback loops might generally lead to spindle centering and that positive feedback loops might generally lead to asymmetrical spindle positioning. Whereas the recent x-ray crystal structures of cytoplasmic dynein (Carter et al., 2011; Kon et al., 2012) have revealed great insights into how the motor walks, they have revealed little about how the motor is locally activated and anchored at the cortex or how GPR-1/LIN-5 switches the motor from a side-on motor to an end-on motor. More attention needs to be focused on the distinction between cortical stiffness and cortical anchoring required in any cortical pulling mechanism. The nonastral spindle positioning mechanisms acting in oocytes of mouse and C. elegans will require a much more detailed understanding of the polarity of cytoplasmic actin filaments and cytoplasmic microtubules.     

Type-3 BRET,an Improved Competition-Based Bioluminescence Resonance Energy Transfer Assay

The heart adjusts its power output to meet specific physiological needs through the coordination of several mechanisms, including force-induced changes in contractility of the molecular motor, the β-cardiac myosin (βCM). Despite its importance in driving and regulating cardiac power output, the effect of force on the contractility of a single βCM has not been measured. Using single molecule optical-trapping techniques, we found that βCM has a two-step working stroke. Forces that resist the power stroke slow the myosin-driven contraction by slowing the rate of ADP release, which is the kinetic step that limits fiber shortening. The kinetic properties of βCM are affected by load, suggesting that the properties of myosin contribute to the force-velocity relationship in intact muscle and play an important role in the regulation of cardiac power output.The cardiac cycle is a tightly regulated process in which the heart generates power during systole and relaxes during diastole. Appropriate power must be generated to effectively pump blood against cardiac afterload. Dysfunction of this cycle has devastating consequences for affected individuals.Cardiac power output is regulated by several feedback mechanisms (e.g., neuronal, hormonal, mechanical) that ultimately lead to changes in the force and power output of the molecular motor, β-cardiac myosin (βCM). In isolated cardiac fibers and cardiomyocytes, loading the muscle during systole slows contraction and alters power output. It is widely believed that this slowing is partially due to force-induced inhibition of myosin ATPase kinetics, similar to the Fenn Effect in skeletal muscle. However, this hypothesis has not been directly tested at the molecular level. Much of our contemporary view of how power is generated in cardiac muscle is due to in vivo and isolated muscle-fiber studies (1). Substantial progress has been made in understanding the actomyosin interactions required for power generation, but resolving the molecular effects of mechanical load on the ATPase properties of βCM in intact muscle has been challenging. Nevertheless, determining the biophysical parameters that define βCM contractility is key to understanding cardiac regulation and the etiology of several muscle diseases (1).In vitro assays using isolated contractile proteins have been central to advancing our understanding of contractility, although most experiments have been conducted at low resisting loads that do not mimic working conditions. Elegant optical trapping experiments have imposed loads on small ensembles of murine α-cardiac myosin at subsaturating [ATP] (2), and these experiments suggest that force slows α-cardiac myosin kinetics. The kinetic properties of α-cardiac myosin are substantially different from βCM, the primary isoform in the adult human myocardium (3). Thus, experiments using βCM must be performed to determine the unitary force-dependent kinetic parameters of this key molecular motor. We used optical trapping to measure the working-stroke displacement and force dependence of actin-detachment kinetics of single porcine βCM molecules at saturating ATP concentrations. These experiments allow direct measurement of the force-velocity relationship for single βCM molecules and reveal the mechanism of how loads regulate βCM-driven power output.Using the three-bead geometry (4) in which an actin filament is strung between two optically-trapped beads and then lowered over a bead that is sparsely coated with purified full-length porcine ventricular βCM, interactions between single βCM molecules and actin were recorded at 10 μM ATP (Fig. 1 A) (5, 6). Ensemble averages of these interactions were constructed to determine the size and kinetics of the working stroke (7, 8). βCM has an average displacement (6.8 ± 0.04 nm) that is similar to previously characterized muscle myosins (9, 10). Similar to skeletal muscle myosin (10), ensemble averages clearly show that the βCM working stroke is composed of two substeps with average displacements of 4.7 ± 0.05 nm and 1.9 ± 0.05 nm (Fig. 1 B). A single exponential function was fit to the rising-phase of the time-forward ensemble averages, yielding a rate (74 ± 2 s⁻¹) for the transition from state 1 to state 2 (Fig. 1 C). This rate is similar to the biochemical rate of ADP release measured for βCM (64 s⁻¹) (3), indicating that this structural transition is associated with the release of ADP. The rate of the rising phase of the time-reversed ensemble averages (22 ± 0.7 s⁻¹) reports the rate of exit from state 2 and is consistent with the biochemical rate of ATP binding and actomyosin detachment at 10 μM ATP (16 s⁻¹) (3) (Fig. 1 C).Open in a separate windowFigure 1(A) Representative data trace showing actomyosin displacements generated by βCM at 10 μM ATP. (Blue lines) Individual binding events. (B) Ensemble averages of the βCM working stroke generated from averaging 1295 binding interactions collected at 10 μM ATP. Single exponential functions were fit to the data (red lines) and the reported errors are the standard errors from the fit. (C) Cartoon showing an idealized actomyosin interaction with the corresponding mechanical and biochemical states. To see this figure in color, go online.To examine actomyosin detachment kinetics under working conditions, a positional feedback optical clamp was used to apply a dynamic load to the myosin, keeping the myosin at an isometric position during its working stroke (11). We measured the effect of force on the actin-attachment duration at 4 mM Mg.ATP to ensure that the rate of ATP binding is not rate-limiting for detachment. Increases in attachment durations are observed as the force on the myosin is increased (Fig. 2 A, inset). Assuming a two-state model (12), we expect the attachment durations to be exponentially distributed at each force with the force-dependent actin detachment rate, k(F), given by (13)k(F)=k0∗e−F·ddetkB∗T,(1)where k₀ is the rate of the primary force-sensitive transition in the absence of force, F is the force on the myosin, d_det is the distance to the transition state (a measurement of force sensitivity), k_B is Boltzmann’s constant, and T is the temperature. Maximum likelihood estimation (MLE) fitting Eq. 1 to the data yields a detachment rate (k₀ = 71 (−1.0/+0.8 s⁻¹)) that is similar to the rate of ADP release measured for βCM (64 s⁻¹) (3) and the rate of the time-forward ensemble averages (74 ± 2 s⁻¹). Thus, the ADP release step (and the accompanying state-1 to state-2 mechanical transition) is force-sensitive (d_det = 0.97 (−0.014/+0.011) nm). The value of d_det indicates that the ADP release step slows with increasing force, but less than some other characterized myosins (14). Using the values determined from the MLE fitting and the measured size of the working stroke, it is possible to calculate a force-velocity relationship for βCM, assuming the rate of ADP release limits actin motility (Fig. 2 A).Open in a separate windowFigure 2(A, Inset) Single molecule actomyosin interactions were collected in the presence of the isometric optical clamp. The scatter plot shows 262 binding events. Attachment durations are exponentially distributed at each force. (A) The detachment rate as a function of force as determined by MLE fitting. (Black line) Best fit; (small gray shaded area) 95% confidence interval. (Right axis) Velocity, calculated by multiplying the displacement of the working stroke by the detachment rate. (B) The calculated mean detachment rate as a function of force. Attachment durations were binned according to the average force experienced by the myosin during the binding event. Error bars were calculated via bootstrapping simulations of each force bin. (Blue line) Expected mean detachment rate based on the MLE fitting and the limited temporal resolution of our experiment (see the Supporting Material for details). (C) Proposed model for how force slows shortening velocity. Force inhibits the mechanical transition associated with ADP release, slowing the rate of actomyosin detachment. To see this figure in color, go online.The MLE fitting of Eq. 1 assumes an exponential distribution of attachment durations at every force. As such, the MLE fitting of the raw data should yield correct values of the parameters k₀ and d_det, despite limitations of the temporal resolution of our experiment (see Supporting Material for detailed discussion of MLE fitting). Frequently, groups report the mean attachment duration as a function of force. However, the mean attachment duration at each force will be overestimated because some shorter binding events cannot be resolved. We provide a method for calculating the expected mean detachment rate based on the parameters determined from the MLE fitting, given the limited temporal resolution of the experiment, and verify the robustness of the MLE fitting (see the Supporting Material). For demonstration purposes only, Fig. 2 B shows that the measured mean detachment rate agrees well with the expected mean detachment rate based on the MLE fitting and the temporal resolution of the experiment. It should be emphasized that the relevant dissociation values are obtained from the MLE fitting in Fig. 2 A (see also Figs. S1–S3).Our data demonstrate that at saturating [ATP], the detachment rate is limited by the ADP release step, which is the same transition that limits fiber shortening velocity (15). We propose that resisting loads slow ADP release and actin detachment by slowing the mechanical transition that accompanies ADP release (Fig. 2 C), thereby reducing the shortening velocity of muscle fibers. Thus, our data demonstrate that the intrinsic force-dependent properties of βCM contribute to the force-velocity relationship in the heart. It is important to note that our proposed mechanism does not rule out additional mechanisms by which force could directly modulate the activity of actomyosin such as force-induced reversal of the power stroke (11) or population of branched pathways (16, 17).Are the loads in our experiments physiologically relevant to contracting muscle? Modeling of the force per cross-bridge generated in isometric soleus muscle, which contains the βCM isoform, suggests a load of 2–4 pN per myosin (18). At these loads, we expect actin-detachment to slow up to threefold. Interestingly, βCM is substantially less force-sensitive than smooth muscle myosin (d_det = 2.7), suggesting that βCM can generate more power (the product of force and velocity) under load.In conclusion, our data show that cardiac power output can be directly modulated by force at the level of single myosin molecules. These data will enable the comparison of how molecular changes, such as light-chain phosphorylation, pharmacological treatments, or mutations associated with cardiomyopathies, affect the ability of the myosin to generate power against the afterload.     

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.     

Big steps toward understanding dynein

Dynein is a microtubule-based molecular motor that is involved in various biological functions, such as axonal transport, mitosis, and cilia/flagella movement. Although dynein was discovered 50 years ago, the progress of dynein research has been slow due to its large size and flexible structure. Recent progress in understanding the force-generating mechanism of dynein using x-ray crystallography, cryo-electron microscopy, and single molecule studies has provided key insight into the structure and mechanism of action of this complex motor protein.It has been 50 years since dynein was discovered and named by Ian Gibbons as a motor protein that drives cilia/flagella bending (Gibbons, 1963; Gibbons and Rowe, 1965). In the mid-1980s, dynein was also found to power retrograde transport in neurons (Paschal and Vallee, 1987). Subsequently, the primary amino acid sequence of the cytoplasmic dynein heavy chain, which contains the motor domain, was determined from the cDNA sequence (Mikami et al., 1993; Zhang et al., 1993). Like other biological motors, such as kinesins and myosins, the amino acid sequence of the dynein motor domain is well conserved. There are 16 putative genes that encode dynein heavy chains in the human genome (Yagi, 2009). Among these is one gene encoding cytoplasmic dynein heavy chain and one encoding retrograde intraflagellar transport dynein heavy chain, while the rest encode for heavy chains of axonemal dyneins. Most of the genes encoding the human dynein heavy chain have a counterpart in Chlamydomonas reinhardtii, which suggests that their functions are conserved from algae to humans.Dynein is unique compared with kinesin and myosin because dynein molecules form large molecular complexes. For example, one axonemal outer arm dynein molecule of C. reinhardtii is composed of three dynein heavy chains, two intermediate chains, and more than ten light chains (King, 2012). Mammalian cytoplasmic dynein consists of two heavy chains and several smaller subunits (Fig. 1 A; Vallee et al., 1988; Allan, 2011). The cargoes of cytoplasmic dynein are various membranous organelles, including lysosomes, endosomes, phagosomes, and the Golgi complex (Hirokawa, 1998). It is likely that one cytoplasmic dynein heavy chain can adapt to diverse cargos and functions by changing its composition.Open in a separate windowFigure 1.Atomic structures of cytoplasmic dynein. (A) Schematic structure of cytoplasmic dynein complex, adapted from Allan (2011). (B) The primary structure of cytoplasmic dynein. (C and D) The atomic model of D. discoideum cytoplasmic dynein motor domain (PDB accession no. 3VKG) overlaid on a microtubule (EMDB-5193; Sui and Downing, 2010) according to the orientation determined by Mizuno et al. (2007) (C) Side view. (D) View from the plus end of microtubule. (E) Schematic domain structure of dynein.Dynein must have a distinct motor mechanism from kinesin and myosin, because it belongs to the AAA+ family of proteins and does not have the conserved amino acid motifs, called the switch regions, present in kinesins, myosins, and guanine nucleotide-binding proteins (Vale, 1996). Therefore, studying dynein is of great interest because it will reveal new design principles of motor proteins. This review will focus on the mechanism of force generation by cytoplasmic and axonemal dynein heavy chains revealed by recent structural and biophysical studies.

Anatomy of dynein

To understand the chemomechanical cycle of dynein based on its molecular structure, it is important to obtain well-diffracting crystals and build accurate atomic models. Recently, Kon and colleagues determined the crystal structures of Dictyostelium discoideum cytoplasmic dynein motor domain, first at 4.5-Å resolution (Kon et al., 2011), and subsequently at 2.8 Å (without the microtubule binding domain) and 3.8-Å (wild type) resolution (Kon et al., 2012). Carter and colleagues also determined the crystal structures of the Saccharomyces cerevisiae (yeast) cytoplasmic dynein motor domain, first at 6-Å resolution (Carter et al., 2011), and later at 3.3–3.7-Å resolution (Schmidt et al., 2012). According to these crystal structures as well as previous EM studies, the overall structure of the dynein heavy chain is divided into four domains: tail, linker, head, and stalk (Fig. 1, B–E). Simply put, each domain carries out one essential function of a motor protein: the tail is the cargo binding domain, the head is the site of ATP hydrolysis, the linker is the mechanical amplifier, and the stalk is the track-binding domain.The tail, which is not part of the motor domain and is absent from crystal structures, is located at the N-terminal ∼1,400 amino acid residues and involved in cargo binding (gray in Fig. 1, B and E). The next ∼550 residues comprise the “linker” (pink in Fig. 1, B–E), which changes its conformation depending on the nucleotide state (Burgess et al., 2003; Kon et al., 2005). This linker domain was first observed by negative staining EM in combination with single particle analysis of dynein c, an isoform of inner arm dynein from C. reinhardtii flagella (Burgess et al., 2003). According to the crystal structures, the linker is made of bundles of α-helices and lies across the AAA+ head domain, forming a 10-nm-long rod-like structure (Fig. 1, C and D). Recent class averaged images of D. discoideum cytoplasmic dynein show that the linker domain is stiff along its entire length when undocked from the head (Roberts et al., 2012). The head (motor) domain of dynein is composed of six AAA+ (ATPase associated with diverse cellular activities) modules (Neuwald et al., 1999; color-coded in Fig. 1, B–E). Although many AAA+ family proteins are a symmetric homohexamer (Ammelburg et al., 2006), the AAA+ domains of dynein are encoded by a single heavy chain gene and form an asymmetric heterohexamer. Among the six AAA+ domains, hydrolysis at the first AAA domain mainly provides the energy for dynein motility (Imamula et al., 2007; Kon et al., 2012). The hexameric ring has two distinct faces: the linker face and the C-terminal face. The linker face is slightly convex and the linker domain lies across this side (Fig. 1 D, left side). The other side of the ring has the C-terminal domain (Fig. 1 D, right side).The stalk domain of dynein was identified as the microtubule-binding domain (MTBD; Gee et al., 1997). It emanates from the C-terminal face of AAA4 and is composed of antiparallel α-helical coiled-coil domain (yellow in Fig. 1, B–E). The tip of the stalk is the actual MTBD. Interestingly, the crystal structures revealed another antiparallel α-helical coiled coil that emerges from AAA5 (orange in Fig. 1, B–E), and this region is called the buttress (Carter et al., 2011) or strut (Kon et al., 2011), which was also observed as the bifurcation of the stalk by negative-staining EM (Burgess et al., 2003; Roberts et al., 2009). The tip of the buttress/strut is in contact with the middle of the stalk and probably works as a mechanical reinforcement of the stalk.

The chemomechanical cycle of dynein

Based on structural and biochemical data, a putative chemomechanical cycle of dynein is outlined in Fig. 2 (A–E). In the no-nucleotide state, dynein is bound to a microtubule through its stalk domain, and its tail region is bound to cargoes (Fig. 2 A). The crystal structures of yeast dynein are considered to be in this no-nucleotide state. When ATP is bound to the AAA+ head, the MTBD quickly detaches from the microtubule (Fig. 2 B; Porter and Johnson, 1983). The ATP binding also induces “hinging” of the linker from the head (Fig. 2 C). According to the biochemical analysis of recombinant D. discoideum dynein (Imamula et al., 2007), the detachment from the microtubule (Fig. 2, A and B) is faster than the later hinging (Fig. 2, B and C). As a result of these two reactions, the head rotates or shifts toward the minus end of the microtubule (for more discussion about “rotate” versus “shift” see the “Dyneins in the axoneme” section) and the MTBD steps forward. The directionality of stepping seems to be mainly determined by the MTBD, because the direction of dynein movement does not change even if the head domain is rotated relative to the microtubule by insertion or deletion of the stalk (Carter et al., 2008). In the presence of ADP and vanadate, dynein is considered to be in this state (Fig. 2 C).Open in a separate windowFigure 2.Presumed chemomechanical cycle and stepping of dynein. (A–E) Chemomechanical cycle of dynein. The pre- and post-power stroke states are also called the primed and unprimed states, respectively. The registries of the stalk coiled coil are denoted as α and β according to Gibbons et al. (2005). (F and G) Processive movement of kinesin (F) and dynein (G). (F) Hand-over-hand movement of kinesin. A step by one head (red) is always followed by the step of another head (green). The stepping of kinesin is on one protofilament of microtubule. (G) Presumed stepping of dynein. The step size varies and the interhead separation can be large. A step by one head (red) is not always flowed by the step of another head (green). (H) A model of strain-based dynein ATPase activation. (G, top) Without strain, the gap between the AAA1 and AAA2 is open and the motor domain cannot hydrolyze ATP. (G, bottom) Under a strain imposed between MTBD and tail (thin black arrows), the gap becomes smaller (thick black arrows) and turns on ATP hydrolysis by dynein.After the MTBD rebinds to the microtubule at the forward site (Fig. 2 D), release of hydrolysis products from the AAA+ head is activated (Holzbaur and Johnson, 1989) and the hinged linker goes back to the straight conformation (Fig. 2 E; Kon et al., 2005). The crystal structure of D. discoideum dynein is considered to be in the state after phosphate release and before ADP release. This straightening of the linker is considered to be the power-generating step and brings the cargo forward relative to the microtubule.

The MTBD of dynein

As outlined in Fig. 2, the nucleotide state of the head domain may control the affinity of the MTBD to the microtubule. Conversely, the binding of the MTBD to the microtubule should activate the ATPase activity of the head domain. This two-way communication is transmitted through the simple ∼17-nm-long α-helical coiled-coil stalk and the buttress/strut, and its structural basis has been a puzzling question.Currently there are three independent MTBD atomic structures in the Protein Data Bank (PDB): One of the crystal structures of the D. discoideum dynein motor domain contains the MTBD (Fig. 3 A), and Carter et al. (2008) crystallized the MTBD of mouse cytoplasmic dynein fused with a seryl tRNA-synthetase domain (Fig. 3 C). The MTBD structure of C. reinhardtii axonemal dynein was solved using nuclear magnetic resonance (PDB accession no. 2RR7; Fig. 3 B). The MTBD is mostly composed of α-helices and the three structures are quite similar to each other within the globular MTBD (Fig. 3). Note that dynein c has an additional insert at the MTBD–microtubule interface (Fig. 3 B, inset), whose function is not yet clear. The three structures start to deviate from the junction between the MTBD and the coiled-coil region of the stalk (Fig. 3, A–C, blue arrowheads). Particularly, one of the stalk α-helix (CC2) in D. discoideum dynein motor domain appears to melt at the junction with the MTBD (Fig. 3 A, red arrowhead). This structural deviation suggests that the stalk coiled coil at the junction is flexible, which is consistent with the observation by EM (Roberts et al., 2009).Open in a separate windowFigure 3.Atomic models of the MTBD of dynein. (A) D. discoideum cytoplasmic dynein (PDB accession no. 3VKH). (B) C. reinhardtii dynein c (PDB accession no. 2RR7). The inset shows the side view, highlighting the dynein c–specific insert. (C) Mouse cytoplasmic dynein (PDB accession no. 3ERR). (D) Mouse cytoplasmic dynein fit to the MTBD–microtubule complex derived from cryo-EM (PDB accession no. 3J1T). All the MTBD structures were aligned using least square fits and color-coded with a gradient from the N to C terminus. CC1, coiled coil helix 1; CC2, coiled coil helix 2. The blue arrowheads points to the junction between MTBD and the stalk, where a well-conserved proline residue (colored pink) is located. In C and D, two residues (isoleucine 3269 and leucine 3417) are shown as spheres. The two residues form hydrophobic contacts in the β-registry (C), whereas they are separated in the α-registry (D) because of the sliding between the two α-helices (blue and red arrows). Conformational changes observed in the mouse dynein MTBD in complex with a microtubule by cryo-EM are shown by black arrows. Note that the cryo-EM density map does not have enough resolution to observe sliding between CC1 and CC2. The sliding was modeled based on targeted molecular dynamics (Redwine et al., 2012).Various mechanisms have been proposed to explain how the affinity between the MTBD and a microtubule is controlled. Gibbons et al. (2005) proposed “the helix-sliding hypothesis” (for review see Cho and Vale, 2012). In brief, this hypothesis proposes that the sliding between two α-helices CC1 and CC2 (Fig. 3, C and D; blue and red arrows) may control the affinity of this domain to a microtubule. When Gibbons’s classification (Gibbons et al., 2005) of the sliding state is applied to the three MTBD structures, the stalk in the D. discoideum dynein motor domain is in the “α-registry” state (not visible in Fig. 3 A because of the melting of CC2), which corresponds to the strong binding state. However, the mouse cytoplasmic and C. reinhardtii axonemal MTBDs have the “β-registry” stalk (Fig. 3 C), which corresponds to the weak binding state.To observe conformational changes induced by the α-registry and/or microtubule binding, Redwine et al. (2012) solved the structure of mouse dynein MTBD in complex with a microtubule at 9.7-Å resolution using cryo-EM and single particle analysis. The MTBD was coupled with seryl tRNA-synthetase to fix the stalk helix in the α-registry. At this resolution, α-helices are visible, and they used molecular dynamics to fit the crystal structure of mouse MTBD (β-registry) to the cryo-EM density map. According to this result, the first helix H1 moves ∼10 Å to a position that avoids a clash with the microtubule (Fig. 3 D, black arrows). This also induces opening of the stalk helix (CC1). Together with mutagenesis and single-molecule motility assays, Redwine et al. (2012) proposed that this new structure represents the strong binding state. Currently, it is not clear why the MTBD structure of D. discoideum dynein motor domain (α-registry, Fig. 3 A) is not similar to the new α-registry mouse dynein MTBD, and this problem needs to be addressed by further studies.

Structures around the first ATP binding site

Another central question about motor proteins is how Ångstrom-scale changes around the nucleotide are amplified to generate steps >8 nm. For dynein, the interface between the first nucleotide-binding pocket and the linker seem to be the key force-generating element (Fig. 4). The crystal structures of dynein give us clues about how nucleotide-induced conformational changes may be transmitted to and amplified by the linker domain.Open in a separate windowFigure 4.Structures around the first ATP binding site. (A) Schematic domain structure of the head domain. Regions contacting the linker domain are colored purple. (B) AAA submodules surrounding the first nucleotide-binding pocket (PDB accession no. 3VKG, chain A). The linker is connected to AAA1 domain by the “N-loop.” To highlight that the two finger-like structures are protruding, the shadow of the atomic structure has been cast on the plane parallel to the head domain. (C) Interaction between the linker and the two finger-like structures. The pink arrowhead points to the hinge-like structure of the linker. The pink numbers indicates the subdomain of the linker.The main ATP catalytic site is located between AAA1 and AAA2 (Fig. 4, A and B). There are four ADP molecules in the D. discoideum dynein crystal structures, but the first ATP binding site alone drives the microtubule-activated ATPase activity, based on biochemical experiments on dyneins whose ATP binding sites were mutated (Kon et al., 2012).One AAA+ module is composed of a large submodule and a small α submodule (Fig. 4 B). The large α/β submodule is located inside of the ring and the small α submodule is located outside. The large submodule bulges toward the linker face, and the overall ring forms a dome-like shape (Fig. 1 D).The main ATP catalytic site is surrounded by three submodules: AAA1 large α/β, AAA1 small α, and AAA2 large α/β (Fig. 4, A and B). Based on the structural changes of other AAA+ proteins (Gai et al., 2004; Suno et al., 2006; Wendler et al., 2012), the gap between AAA1 and AAA2 modules is expected to open and close during the ATPase cycle.In fact, the size of the gap varies among the dynein crystal structures. The crystal structures of yeast dyneins show a larger gap between AAA1 and AAA2, which might be the reason why no nucleotide was found in the binding pocket. Although Schmidt et al. (2012) soaked the crystals in a high concentration of various nucleotides (up to 25 mM of ATP), no electron densities corresponding to the nucleotide were observed at the first ATP binding site. Among dynein crystal structures, one of D. discoideum dynein (PDB accession no. 3VKH, chain A) has the smallest gap, but it is still considered to be in an “open state” because the arginine finger in the AAA2 module (Fig. 4 B, red) is far from the phosphates of ADP. Because the arginine finger is essential for ATP hydrolysis in other AAA+ proteins (Ogura et al., 2004), the gap is expected to close and the arginine finger would stabilize the negative charge during the transition state of ATP hydrolysis.The presumed open/close conformational change between AAA1 and AAA2 would result in the movement of two “finger-like” structures protruding from the AAA2 large α/β submodule (Fig. 4 B). The two finger-like structures are composed of the H2 insert β-hairpin and preSensor I (PS-I) insert. In D. discoideum dynein crystal structure, the two finger-like structures are in contact with the “hinge-like cleft” of the linker (Fig. 4 C, pink arrowhead). The hinge-like cleft is one of the thinnest parts of the linker, where only one α-helix is connecting between the linker subdomains 2 and 3.In the yeast crystal structures, which have wider gaps between AAA1 and AAA2, the two finger-like structures are not in direct contact with the linker and separated by 18 Å. Instead, the N-terminal region of the linker is in contact with the AAA5 domain (Fig. 4 A). To test the functional role of the linker–AAA5 interaction, Schmidt et al. (2012) mutated a residue involved in the interaction (Phe3446) and found that the mutation resulted in severe motility defects, showing strong microtubule binding and impaired ATPase activities. In D. discoideum dynein crystals, there is no direct interaction between AAA5 and the linker, which suggests that the gap between AAA1 and AAA2 may influence the interaction between the head and linker domain. The contact between the linker and AAA5 may also influence the gap around AAA5, because the gap between AAA5 and AAA6 is large in yeast dynein crystal, whereas the one between AAA4 and AAA5 is large in D. discoideum dynein.The movement of two finger-like structures would induce remodeling of the linker. According to the recent cryo-EM 3D reconstructions of cytoplasmic dynein and axonemal dynein c (Roberts et al., 2012), the linker is visible across the head and there is a large gap between AAA1 and AAA2 in the no-nucleotide state. This linker structure is considered to be the “straight” state (Fig. 2, A and E). In the presence of ADP vanadate, the gap between AAA1 and AAA2 is closed and the N-terminal region of linker is near AAA3, which corresponds to the pre-power stroke “hinged” state (Fig. 2, C and D). The transition from the hinged state to the straight state of the linker is considered to be the force-generating step of dynein.

Processivity of dynein

As the structure of dynein is different from other motor proteins, dynein’s stepping mechanism is also distinct. Both dynein and kinesin are microtubule-based motors and move processively. Based on the single molecule tracking experiment with nanometer accuracy (Yildiz et al., 2004), it is widely accepted that kinesin moves processively by using its two motor domain alternately, called the “hand-over-hand” mechanism. To test whether dynein uses a similar mechanism to kinesin or not, recently Qiu et al. (2012) and DeWitt et al. (2012) applied similar single-molecule approaches to dynein.To observe the stepping, the two head domains of yeast recombinant cytoplasmic dynein were labeled with different colors and the movement of two head domains was tracked simultaneously. If dynein walks by the hand-over-hand mechanism, the step size would be 16 nm and the stepping of one head domain would always be followed by the stepping of another head domain (alternating pattern), and the trailing head would always take a step (Fig. 2 F). Contrary to this prediction, both groups found that the stepping of the head domains is not coordinated when the two head domains are close together. These observations indicated that the chances of a leading or trailing head domain stepping are not significantly different (Fig. 2 G; DeWitt et al., 2012; Qiu et al., 2012).This stepping pattern predicts that the distance between the head domains can be long. In fact, the distance between the two head domains is on average ∼18 ± 11 nm (Qiu et al., 2012) or 28.4 ± 10.7 nm (DeWitt et al., 2012), and as large as ∼50 nm (DeWitt et al., 2012). When the two head domains are separated, there is a tendency where stepping of the trailing head is preferred over that of the forward head.In addition, even though the recombinant cytoplasmic dynein is a homodimer, the two heavy chains do not function equally. While walking along the microtubule, the leading head tends to walk on the right side, whereas the trailing head walks on the left side (DeWitt et al., 2012; Qiu et al., 2012). This arrangement suggests that the stepping mechanism is different between the two heads. In fact, when one of the two dynein heavy chains is mutated to abolish the ATPase activity at AAA1, the heterodimeric dynein still moves processively (DeWitt et al., 2012), with the AAA1-mutated dynein heavy chain remaining mostly in the trailing position. This result clearly demonstrates that allosteric communication between the two AAA1 domains is not required for processivity of dynein. It is likely that the mutated head acts as a tether to the microtubule, as it is known that wild-type dynein can step processively along microtubules under external load even in the absence of ATP (Gennerich et al., 2007).These results collectively show that dynein moves by a different mechanism from kinesin. It is likely that the long stalk and tail allow dynein to move in a more flexible manner.

Dyneins in the axoneme

As mentioned in the introduction, >10 dyneins work in motile flagella and cilia. The core of flagella and cilia is the axoneme, which is typically made of nine outer doublet microtubules and two central pair microtubules (“9 + 2,” Fig. 5 A). The axonemes are found in various eukaryotic cells ranging from the single-cell algae C. reinhardtii to human. Recent extensive cryo-electron tomography (cryo-ET) in combination with genetics revealed the highly organized and complex structures of axonemes that are potentially important for regulating dynein activities (Fig. 5, C and D; Nicastro et al., 2006; Bui et al., 2008, 2009, 2012; Heuser et al., 2009, 2012; Movassagh et al., 2010; Lin et al., 2012; Carbajal-González et al., 2013; Yamamoto et al., 2013).Open in a separate windowFigure 5.Arrangement of axonemal dyneins. (A) The schematic structure of the motile 9 + 2 axoneme, viewed from the base of flagella. (B) Quasi-planar asymmetric movement of the 9 + 2 axoneme typically observed in trachea cilia or in C. reinhardtii flagella. (C and D) 3D structure of a 96-nm repeat of doublet microtubules in the distal/central region of C. reinhardtii flagella (EMDB-2132; Bui et al., 2012). N-DRC, the nexin-dynein regulatory complex; ICLC, intermediate chain/light chain complex. Inner arm dynein subspecies are labeled according to Bui et al. (2012) and Lin et al. (2012). To avoid the confusion with the linker domain of dynein, the structures connecting between outer and inner arm dyneins are labeled as “connecters,” which are normally called “linkers.” Putative ATP binding sites of outer arm dynein determined by biotin-ADP (Oda et al., 2013) are indicated by orange circles. The atomic structure of cytoplasmic dynein is placed into the β-heavy chain of outer arm dynein and its enlarged view is shown in the inset. (D) Two doublet microtubules, viewed from the base of flagella.The basic mechanochemical cycles of axonemal dyneins are believed to be shared with cytoplasmic dynein. Dynein c is an inner arm dynein of C. reinhardtii and used extensively to investigate the conformational changes of dynein, as shown in Fig. 2 (A–E), by combining EM and single-particle analysis (Burgess et al., 2003; Roberts et al., 2012). Structural changes of axonemal dyneins complexed with microtubules are also observed by quick-freeze and deep-etch EM (Goodenough and Heuser, 1982; Burgess, 1995), cryo-EM (Oda et al., 2007), negative-staining EM (Ueno et al., 2008), and cryo-ET (Movassagh et al., 2010). According to these studies, the AAA+ head domains are constrained near the A-tubule in the no-nucleotide state. In the presence of nucleotide, the head domains move closer to the B-tubule and/or the minus end of microtubule, and their appearance becomes heterogeneous, which is consistent with the observation of isolated dynein c that shows greater flexibility between tail and stalk in the ADP/vanadate state (Burgess et al., 2003).One of the controversies about the structural changes of axonemal dyneins is whether their stepping involves “rotation” or “shift” of the head (Fig. 2, B to D). The stalk angle relative to the microtubule seems to be a constant ∼60° irrespective of the nucleotide state (Ueno et al., 2008; Movassagh et al., 2010). This angle is similar to the angle obtained from cryo-EM study of the MTBD–microtubule complex (Redwine et al., 2012). Based on these observations, Ueno et al. (2008) and Movassagh et al. (2010) hypothesize that the “shift” of the head pulls the B-microtubule toward the distal end. However, Roberts et al. (2012) propose that the “rotation” of head and stalk is involved in the stepping based on the docking of dynein c head into an averaged flagella tomogram obtained by Movassagh et al. (2010). This issue needs to be resolved by more reliable and high-resolution data, but these two models may not be mutually exclusive. For example, averaged tomograms may be biased toward the microtubule-bound stalk because tomograms are aligned using microtubules.To interpret these structural changes of axonemal dyneins, docking atomic models of dynein is necessary. According to Roberts et al. (2012), the linker face of inner arm dynein c is oriented outside of axoneme (Fig. 5 D). For outer arm dyneins, we used cryo-EM in combination with biotin-ADP-streptavidin labeling and showed that the ATP binding site, most likely AAA1, is on the left side of the AAA+ head (Fig. 5 C; Oda et al. (2013)). Assuming that the stalks extend out of the plane toward the viewer, the linker face of outer arm dynein is oriented outside of axoneme (Fig. 5 C, inset; and Fig. 5 D). If it were the opposite, the AAA1 would be located on the right side of the AAA+ head. In summary, both inner and outer arm dynein seem to have the same arrangement, with their linker face oriented outside of the axoneme (Fig. 5 D).A unique characteristic of axonemal dyneins is that these dyneins are under precise temporal and spatial control. To generate a planer beating motion (Fig. 5 B), dyneins should be asymmetrically controlled, because the dyneins located on doublets 2–4 drive the effective stroke, whereas the ones on doublets 6–8 drive the recovery stroke (Fig. 5 A). Based on the cryo-ET observation of axonemes, Nicastro et al. (2006) proposed that “linkers” between dyneins provide hard-wiring to coordinate motor activities. Because the linkers in axonemes are distinct structures from the linker domain of dynein, for clarity, here we call them “connecters.” According to the recent cryo-ET of proximal region of C. reinhardtii flagella (Bui et al., 2012), there are in fact asymmetries among nine doublets that are localized to the connecters between outer and inner arm dynein, called the outer-inner dynein (OID) connecters (Fig. 5, A and C). Recently we identified that the intermediate chain 2 (IC2) of outer arm dynein is a part of the OID connecters, and a mutation of the N-terminal region of IC2 affects functions of both outer and inner arm dyneins (Oda et al., 2013), which supports the idea that the connecters between dyneins are involved in axonemal dynein regulation.

Closing remarks

Thanks to the crystal structures, we can now design and interpret experiments such as single molecule assays and EM based on the atomic models of dynein. Our understanding of the molecular mechanism and cellular functions of dyneins will be significantly advanced by these experiments in the near future.One important direction of dynein research is to understand the motor mechanisms closer to the in vivo state. For example, the step sizes of cytoplasmic dynein purified from porcine brain is ∼8 nm independent of load (Toba et al., 2006). This result suggests that intermediate and light chain bound to the dynein heavy chain may modulate the motor activity of dynein. To address such questions, Trokter et al. (2012) reconstituted human cytoplasmic dynein complex from recombinant proteins, although the reconstituted dynein did not show robust processive movement. Further studies are required to understand the movement of cytoplasmic dynein. Similarly, axonemal dyneins should also be studied using mutations in a specific gene that does not affect the overall flagella structure, rather than depending on null mutants that cause the loss of large protein complexes.Detailed full chemomechanical cycle of dynein and its regulation are of great importance. Currently, open/closed states of the gap between AAA1 and AAA2 are not clearly correlated with the chemomechanical cycle of dynein. Soaking dynein crystal with nucleotides showed that the presence of ATP alone is not sufficient to close the gap, at least in the crystal (Schmidt et al., 2012). This result suggests that other factors such as a conformational change of the linker are required. For other motors, ATP hydrolysis is an irreversible chemical step, which is often “gated” by strain. In the case of kinesin, ATP is hydrolyzed by a motor domain only when a forward strain is applied by the other motor domain through the neck linker (Cross, 2004; Kikkawa, 2008). A similar strain-based gating mechanism may play important roles in controlling the dynein ATPase. Upon MTBD binding to the forward binding site, a strain between MTBD and tail would be applied to the dynein molecule. The Y-shaped stalk and strut/buttress under the strain would force the head domain to close the gap between AAA4 and AAA5 (Fig. 2 H). Similarly, the linker under the strain would be hooked onto the two finger-like structures and close the gap between AAA1 and AAA2 (Fig. 2 H). The gap closure then triggers ATP hydrolysis by dynein. This strain-based gating of dynein is consistent with the observation that the rate of nonadvancing backward steps, which would depend on ATP hydrolysis, is increased by load applied to dynein (Gennerich et al., 2007). To explain cilia and flagella movement, the geometric clutch hypothesis has been proposed (Lindemann, 2007), which contends that the forces transverse (t-force) to the axonemal axis act on the dynein to regulate dynein activities. In the axoneme, dynein itself can be the sensor of the t-force by the strain-based gating mechanism. Further experiments are required to test this idea, but the strain-based gating could be a shared property of biological motors.     

Hypoxia-cultured human adipose-derived mesenchymal stem cells are non-oncogenic and have enhanced viability,motility, and tropism to brain cancer

Adult human adipose-derived mesenchymal stem cells (hAMSCs) are multipotent cells, which are abundant, easily collected, and bypass the ethical concerns that plague embryonic stem cells. Their utility and accessibility have led to the rapid development of clinical investigations to explore their autologous and allogeneic cellular-based regenerative potential, tissue preservation capabilities, anti-inflammatory properties, and anticancer properties, among others. hAMSCs are typically cultured under ambient conditions with 21% oxygen. However, physiologically, hAMSCs exist in an environment of much lower oxygen tension. Furthermore, hAMSCs cultured in standard conditions have shown limited proliferative and migratory capabilities, as well as limited viability. This study investigated the effects hypoxic culture conditions have on primary intraoperatively derived hAMSCs. hAMSCs cultured under hypoxia (hAMSCs-H) remained multipotent, capable of differentiation into osteogenic, chondrogenic, and adipogenic lineages. In addition, hAMSCs-H grew faster and exhibited less cell death. Furthermore, hAMSCs-H had greater motility than normoxia-cultured hAMSCs and exhibited greater homing ability to glioblastoma (GBM) derived from brain tumor-initiating cells from our patients in vitro and in vivo. Importantly, hAMSCs-H did not transform into tumor-associated fibroblasts in vitro and were not tumorigenic in vivo. Rather, hAMSCs-H promoted the differentiation of brain cancer cells in vitro and in vivo. These findings suggest an alternative culturing technique that can enhance the function of hAMSCs, which may be necessary for their use in the treatment of various pathologies including stroke, myocardial infarction, amyotrophic lateral sclerosis, and GBM.Mesenchymal stem cells (MSCs) are multipotent cells, isolated from the bone marrow, adipose tissue, and muscle, among others. They are clonally expansive, with the capacity to differentiate into osteocytes, adipocytes, and chondrocytes.^{1, 2} MSCs are widely studied for their regenerative potential, tissue preservation capabilities, anti-inflammatory properties, and anticancer therapeutic potential.^{3, 4, 5} MSCs can serve as vehicles for delivering effective targeted therapy to primary brain cancer and metastatic cancer.^{6, 7, 8}Notwithstanding aggressive treatment of primary brain cancer (glioblastoma (GBM)) with surgical resection, chemotherapy, and radiotherapy, the median survival following diagnosis is 14.6 months.^{9, 10, 11, 12, 13, 14, 15} GBM-targeted therapy using neural stem cells and MSCs as vehicles for therapeutic agents is a promising strategy.¹⁶ MSCs seem to be the ideal stem cells, as they are autologous, easily collected, and easily re-implanted.^{17, 18} The most commonly used MSCs are bone marrow-derived MSCs (BM-MSCs) and human adipose-derived MSCs (hAMSCs). Compared with BM-MSCs, hAMSCs are easier to obtain.^{19, 20}Despite the potential utility of hAMSCs, their use is hampered by their low concentration within tissues.^{21, 22} Thus, in vitro expansion of hAMSCs is necessary. Compared with BM-MSCs, hAMSCs are more genetically and morphologically stable in long-term culture.^{19, 20} Nevertheless, current culturing conditions for both BM-MSCs and hAMSCs show a progressive decrease in viability and proliferative ability, and an increase in senescence ratio for these stem cells with time.^{23, 24, 25, 26, 27, 28, 29} Typically, hAMSCs are cultured under ambient conditions with 21% oxygen in vitro.³⁰ However, physiologically, hAMSCs exist at much lower oxygen tensions, between 1 and 14%.^{31, 32} As a result of the limitations of culturing under normoxia, we investigated the effect of hypoxia on intraoperatively obtained hAMSCs by assessing proliferation, survival, differentiation, tumor formation, tumor tropism, and migration in vitro and in vivo in a rodent model with a human brain cancer. hAMSCs have been reported to transform into tumor-associated fibroblasts (TAFs), which can potentially support tumor growth and promote malignant phenotypes.^{33, 34} Yet, no studies have reported on the changes that may occur in hypoxia-cultured hAMSCs after they are exposed to brain cancer, both in vitro and in vivo. An understanding of the effects of hypoxia on hAMSCs³⁵ is critical for its potential therapeutic applications including in the treatment of brain tumors, stroke, neuro-degenerative diseases such as multiple sclerosis, and dementia (Figure 1a).Open in a separate windowFigure 1Primary human adipose-derived cells cultured in hypoxia (hAMSCs-H) and normoxia (hAMSCs-N) are both MSCs but normoxia-cultured cells show increased signs of senescence, such as increased area and elongated morphology, compared with hypoxia-cultured cells. (a) hAMSCs were isolated from human fat tissue and cultured in hypoxic (1.5% oxygen) or normoxic (21% oxygen) conditions in vitro. The viability, mobility, tumor tropism, safety, and tumorigenic potential were subsequently compared in vitro and in vivo. (b) Differentiation assay. hAMSCs were cultured in control media and differentiation media for 3 weeks, 10 days after the second passage. Three different stains were performed to assess differentiation capabilities (scale bar, 100 μm). (c) Flow cytometric analysis was performed to confirm the absence of CD31-, CD34-, and CD45-positive cells in both cell cultures. In addition, primary hAMSC cultures expressed high levels of CD73, CD90, and CD105, both in hypoxic and normoxic culture conditions at day 10 after passage 2. (d) Representative images of cell morphologies of hAMSCs on 2D surface (scale bar, 200 μm). (e) Schematic of 3D-nanopatterned surface used to assess morphology and motility. (f) Images of cell morphologies of hAMSCs on 3D-nanopatterned surface (scale bar, 200 μm). (g–j) The length, width, area, and length-to-width ratio were measured and compared after cell aligned on the nanopattern surface. Error bars represent S.E.M. *P<0.05, **P<0.01, N.S., not significant     

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.     

Mitochondrial Protein Synthesis,Import, and Assembly

The mitochondrion is arguably the most complex organelle in the budding yeast cell cytoplasm. It is essential for viability as well as respiratory growth. Its innermost aqueous compartment, the matrix, is bounded by the highly structured inner membrane, which in turn is bounded by the intermembrane space and the outer membrane. Approximately 1000 proteins are present in these organelles, of which eight major constituents are coded and synthesized in the matrix. The import of mitochondrial proteins synthesized in the cytoplasm, and their direction to the correct soluble compartments, correct membranes, and correct membrane surfaces/topologies, involves multiple pathways and macromolecular machines. The targeting of some, but not all, cytoplasmically synthesized mitochondrial proteins begins with translation of messenger RNAs localized to the organelle. Most proteins then pass through the translocase of the outer membrane to the intermembrane space, where divergent pathways sort them to the outer membrane, inner membrane, and matrix or trap them in the intermembrane space. Roughly 25% of mitochondrial proteins participate in maintenance or expression of the organellar genome at the inner surface of the inner membrane, providing 7 membrane proteins whose synthesis nucleates the assembly of three respiratory complexes.TO think about how mitochondrial proteins are synthesized, imported, and assembled, it is useful to have a clear picture of the organellar structures that they, along with membrane lipids, compose and the functions that they carry out. As almost every schoolchild learns, mitochondria carry out oxidative phosphorylation, the controlled burning of nutrients coupled to ATP synthesis. Since Saccharomyces cerevisiae prefers to ferment sugars, respiration is a dispensable function and nonrespiring mutants are viable [although they cannot undergo meiosis (Jambhekar and Amon 2008)]. However, mitochondria themselves are not dispensable. A substantial fraction of intermediary metabolism occurs in mitochondria (Strathern et al. 1982), and at least one of these pathways, iron–sulfur cluster assembly, is essential for growth (Kispal et al. 2005). Thus, any mutation that prevents the biogenesis of mitochondria by, for example, preventing the import of protein constituents from the cytoplasm, is lethal (Baker and Schatz 1991).The mitochondria of S. cerevisiae are tubular structures at the cell cortex. While the number of distinct compartments can range from 1 to ∼50 depending upon conditions (Stevens 1981; Pon and Schatz 1991), continual fusion and fission events among them effectively form a single dynamic network (Nunnari et al. 1997). The outer membrane surrounds the tubules. The inner membrane has a boundary domain closely juxtaposed beneath the outer membrane and cristae domains that project internally from the boundary into the matrix (Figure 1A). The matrix is the aqueous compartment surrounded by the inner membrane. The aqueous intermembrane space lies between the membranes and is continuous with the space within cristae.Open in a separate windowFigure 1 Overview of mitochondrial structure in yeast. (A) Schematic of compartments comprising mitochondrial tubules. The outer membrane surrounds the organelle. The inner membrane surrounds the matrix and consists of two domains, the inner boundary membrane and the cristae membranes, which are joined at cristae junctions. The intermembrane space lies between the outer membrane and inner membrane. (B) Electron tomograph image of a highly contracted yeast mitochondrion observed en face (a) with the outer membrane (red) and (b) without the outer membrane. Reprinted by permission from John Wiley & Sons from Mannella et al. (2001).Inner membrane cristae are often depicted as baffles emanating from the boundary domain. However, electron tomography of mitochondria from several species, including yeast, shows that cristae actually emanate from the boundary membrane as narrow tubular structures at sites termed “crista junctions” and expand as they project into the matrix (Frey and Mannella 2000; Mannella et al. 2001) (Figure 1B). It seems clear that the boundary and cristae domains of the inner membrane have distinct compositions with respect to the respiratory complexes that are embedded preferentially in the cristae membrane domains, as well as other components (Vogel et al. 2006; Wurm and Jakobs 2006; Rabl et al. 2009; Suppanz et al. 2009; Zick et al. 2009; Davies et al. 2011).The outer and inner boundary membranes are connected at multiple contact sites, at least some of which are involved in protein translocation and may be transient (Pon and Schatz 1991). In addition, there appear to be firm contact sites, not directly involved with protein translocation, preferentially colocalized with crista junctions (Harner et al. 2011a).Overall, there appear to be ∼1000 distinct proteins in yeast mitochondria (Premsler et al. 2009). One series of proteomic studies on highly purified organelles identified 851 proteins thought to represent 85% of the total number of species (Sickmann et al. 2003; Reinders et al. 2006; Zahedi et al. 2006). Another study identified an additional 209 candidates (Prokisch et al. 2004). A computationally driven search for candidates involved in yeast mitochondrial function, coupled with experiments to assay respiratory function and maintenance of mitochondrial DNA (mtDNA), identified 109 novel candidates, although many of these may not be mitochondrial per se (Hess et al. 2009). Taking the boundary and cristae domains together, the inner membrane is the most protein-rich mitochondrial compartment, followed by the matrix (Daum et al. 1982).Only eight of the yeast mitochondrial proteins detected in proteomic studies are encoded by mtDNA and synthesized within the organelle. They are hydrophobic subunits of respiratory complexes III (bc₁ complex or ubiquinol-cytochrome c reductase), IV (cytochrome c oxidase), and V (ATP synthase), as well as a hydrophilic mitochondrial small subunit ribosomal protein. The remaining ∼99% of yeast mitochondrial proteins are encoded by nuclear genes, synthesized in cytoplasmic ribosomes, and imported into the organelle.An overview of known nuclearly encoded mitochondrial protein functions (Figure 2) reveals that ∼25% of them are involved directly in genome maintenance and expression of the eight major mitochondrial genes (Schmidt et al. 2010). The functions of ∼20% of the proteins are not known. Fifteen percent are involved in the well-known processes of energy metabolism. Protein translocation, folding, and turnover functions occupy ∼10% of mitochondrial proteins.Open in a separate windowFigure 2 Classification of identified mitochondrial proteins according to function. Reprinted by permission from Nature Publishing Group from Schmidt et al. (2010).The following discussion reviews our understanding of the biogenesis of mitochondria starting on the outside, the cytoplasm, and working inward through the mitochondrial compartments.     

Voltage Sensor Gating Charge Transfer in a hERG Potassium Channel?Model

Relaxation of a hERG K⁺ channel model during molecular-dynamics simulation in a hydrated POPC bilayer was accompanied by transitions of an arginine gating charge across a charge transfer center in two voltage sensor domains. Inspection of the passage of arginine side chains across the charge transfer center suggests that the unique hydration properties of the arginine guanidine cation facilitates charge transfer during voltage sensor responses to changes in membrane potential, and underlies the preference of Arg over Lys as a mobile charge carrier in voltage-sensitive ion channels.The response of voltage-sensitive ion channels to changes in membrane potential is mediated by voltage sensor domains (VSD) containing a transmembrane helical segment (S4) with a repeating motif of positively charged and hydrophobic amino acids (Fig. 1) (1,2). Changes in membrane potential drive the S4 helix through the membrane plane with the charged side chains (largely arginine) on S4 swapping Glu/Asp carboxylate partners that lie on less mobile elements of the VSD (2). Movement of S4 is coupled to the ion-conducting pore to transmit changes in membrane potential to channel gating (3).Open in a separate windowFigure 1Structures of the VSD of membrane domains before MD in a POPC bilayer. The S2 (pink) and S4 (blue) helices of the VSD of the hERG model (A) and Kv1.2/2.1 chimera structure (B) are highlighted. (C) Sequence alignment of S2 and S4 among homologous voltage-sensitive K⁺ channels.The VSD charge-pairing motif of K⁺ and Na⁺ channels is best represented in VSD states at zero membrane potential (S4 helix up) for which crystal structures exist for Kv1.2 (4), Kv1.2/2.1 chimera (5), and Na_v channels (6,7). In these states, positively charged residues on the intra- and extracellular sections of the S4 helix are separated by a hydrophobic charge-transfer center (CTC) (1) or plug (8) containing a highly conserved Phe residue (Fig. 1). This plug restricts water incursion across the VSD, focusing the electric field across a narrow region near the bilayer center. In voltage-driven transitions between S4 down- and up-states, positively charged S4 side chains move across the CTC.The ether-à-go-go (eag) and eag-related family of voltage-sensitive K⁺ channels likely share similar charge pairing interactions with VSDs in other channels (9,10). However, eag VSDs contain an extra negative charge on S2 (underlined in Fig. 1 C) so that in hERG, Asp residues (D460 and D466) lie approximately one helical turn above and below the conserved charge-transfer center Phe (F463) (Fig. 1). This eag-specific motif might be expected to facilitate transfer of Arg side chains through the CTC and to stabilize the voltage sensor (VS) in the up state. We recently described an open state (VS-up) hERG model built on the crystal structure template of the Kv1.2/2.1 chimera and molecular-dynamics (MD) simulation of this model in a hydrated POPC bilayer (11). We have inspected an extended version of this simulation and identified transitions of a gating charge into the CTC despite the absence of a membrane potential change. These transitions are absent in equivalent MD simulations of the chimera structure in a POPC bilayer.Fig. 1 shows a single VS from starting structures of the hERG model and the chimera structure in a hydrated POPC bilayer, after restrained MD to anneal the protein-lipid interface (see Methods in the Supporting Material). Because the hERG model is constructed on the chimera structure according to the alignment in Fig. 1 the pattern of pairing between S4 charges and acidic VS side chains is equivalent in the hERG model and chimera structure.The arrangement of charge-paired side chains remains constant during MD in all subunits of the chimera (e.g., Fig. 2 E and see Fig. S2 in the Supporting Material). However, in two subunits of the hERG model the R534 side chain moves toward the extracellular side of the bilayer, sliding into the CTC to form a charge interaction with the extra Asp residue (D460 in hERG) that lies just above F463 (Fig. 2, A–C). This transition is facilitated by changes in side-chain rotamers of R534 and F463 as the planar Arg guanidine group rotates past the F463 ring, and the availability of D460 as a counterion for the R534 guanidine (Fig. 2). Movement of an Arg guanidine past the Phe side chain of the CTC is similar to that described in steered MD of an isolated VS domain (12).Open in a separate windowFigure 2Movement of the R534 side chain across the CTC in chain a of the hERG model simulation (A). Similar transitions are observed in chains a and b (panels B and C), but not chains c (D) or d (not shown), where the R534 side chain remains close to D466. In all subunits of the Kv1.2/2.1 chimera simulation, charge pairing of the starting structure (Fig. 1B) was maintained throughout (e.g., panel E and see Fig. S2 in the Supporting Material). (Black and blue lines) Distances from the Arg CZ or Lys ε atom to the two O atoms, respectively, of Asp or Glu.Mason et al. (13) have shown, using neutron scattering, that the low charge density guanidine cation (Gdm⁺) corresponding to the Arg side chain is poorly hydrated above and below the molecular plane. This property may underlie the universal preference for Arg (over Lys) in voltage sensor charge transfer. Although the poorly-hydrated surfaces of Gdm⁺ interact favorably with nonpolar (especially planar) surfaces (14,15), Gdm⁺ retains in-plane hydrogen bonding (13). In the transition of R534 across the CTC, in-plane solvation of the guanidine side chain is provided initially by D466, D501, and water molecules below the CTC, and during and after the transition by D501 and D460 side chains and waters above the CTC (Fig. 3, A and B). Complete transfer of the R534 side chain across the CTC was not observed, but would be expected to involve movement of the guanidine group away from H-bonding distance with D501.Open in a separate windowFigure 3In-plane solvation of R534 guanidine in the charge transfer center during the hERG model MD (A). (Dotted lines) H-bond distances of <2.5 Å. The right-hand group consists of top-down (B) and end-on (C) views of the distribution of oxygen atoms around the side chain of hERG R534 at 20-ns intervals during MD (subunit a). (D) End-on view of equivalent atom distributions around the K302 side chain during the Kv1.2/2.1 chimera MD (subunit c). (Red spheres, water O; pink, Asp OD1 and OD2; purple:, Glu OE1 and OE2.)The atom distribution around the R534 side chain during MD (Fig. 3, B and C) conforms to the experimental Gdm⁺ hydration structure (13), with H-bonding to waters and side-chain Asp O atoms exclusively in the guanidine plane. The passage of Gdm⁺ through the CTC is facilitated by the hydrophobic nature of Gdm⁺ above and below the molecular plane (13), which allows interaction with the nonpolar groups (especially F463) in the CTC (Fig. 3 A and see Fig. S3). This contrasts with the solvation properties of the Lys amino group (e.g., K302 of the Kv1.2/2.1 chimera (Fig. 1), which has a spherical distribution of H-bonding and charge-neutralizing oxygen atoms (Fig. 3 D and see Fig. S4).To further test these interpretations, we ran additional MD simulations of the isolated hERG VS domain model and an R534K mutant in a hydrated POPC bilayer. Again, the R534 side chain entered the CTC in the wild-type model simulation whereas the K534 side chain did not (see Fig. S5). Inspection of the atom distributions in Fig. 3 D (and see Fig. S4) indicates that the pocket below the conserved Phe of the CTC is particularly favorable for a Lys side chain, with waters and acidic side chains that satisfy the spherical solvation requirements of the terminal amino group, and nonpolar side chains that interact with the aliphatic part of the side chain.The occurrence of transitions of the R534 side chain through the CTC in the hERG model, in the absence of a change in membrane potential, indicates a relaxation from a less-stable starting structure. However, the path of the R534 side chain provides useful molecular-level insight into the nature of charge transfer in voltage sensors. How do these observations accord with broader evidence of charge transfer in voltage-sensitive channels in general, and hERG in particular? Studies with fluorinated analogs of aromatic side chains equivalent to F463 of hERG or F233 of the chimera indicate the absence of a significant role for cation-π interactions involving the CTC aromatic group in K⁺ and Na_v channels, although a planar side chain is preferred in some cases (1,16). In hERG, F463 can be replaced by M, L, or V with small effects on channel gating (17), indicating that the hERG CTC requires only a bulky nonpolar side chain to seal the hydrophobic center of the VS and allow passage of the Arg side chain through the CTC. Both absence of requirement for cation-π interactions, and accommodation of nonplanar hydrophobic side chains in a functional hERG CTC, are broadly consistent with the interpretation that it is the poorly-hydrated nature of the Arg guanidine group above and below the molecular plane (together with its tenacious proton affinity (18)) that governs its role in carrying gating charge in voltage sensors.While the simulations suggest that R534 may interact with D460 in the open channel state, the possibility that the extra carboxylate side chain above the CTC might facilitate gating charge transfer is seemingly inconsistent with the slow activation of hERG, although hERG D460C does activate even more slowly than the WT channel (9). However, S4 movement in hERG occurs in advance of channel opening (19), and slow gating is partly mediated by interactions involving hERG cytoplasmic domains (20); thus, slow S4 movement may not be an inherent property of the hERG voltage sensor. Recent studies show that when hERG gating is studied at very low [Ca²⁺] (50 μM) and low [H⁺] (pH 8.0), the channel is strongly sensitized in the direction of the open state; this effect is reduced in hERG D460C (and hERG D509C) (10). These observations support a role for the extra hERG Asp residues in binding Ca²⁺ (and H⁺) (10), allowing the channel to be allosterically responsive to changes in pH and [Ca²⁺]. A true comparison of a hERG model with experimental channel gating might involve studies on a channel lacking cytoplasmic domains that modulate gating, and using conditions (high pH and low [Ca²⁺]) that leave the eag-specific Asp residues unoccupied. This could reveal the inherent current-voltage relationships and kinetics of the hERG voltage sensor.     

Novel Growth Regime of MDCK II Model Tissues on Soft Substrates

It is well established that MDCK II cells grow in circular colonies that densify until contact inhibition takes place. Here, we show that this behavior is only typical for colonies developing on hard substrates and report a new growth phase of MDCK II cells on soft gels. At the onset, the new phase is characterized by small, three-dimensional droplets of cells attached to the substrate. When the contact area between the agglomerate and the substrate becomes sufficiently large, a very dense monolayer nucleates in the center of the colony. This monolayer, surrounded by a belt of three-dimensionally packed cells, has a well-defined structure, independent of time and cluster size, as well as a density that is twice the steady-state density found on hard substrates. To release stress in such dense packing, extrusions of viable cells take place several days after seeding. The extruded cells create second-generation clusters, as evidenced by an archipelago of aggregates found in a vicinity of mother colonies, which points to a mechanically regulated migratory behavior.Studying the growth of cell colonies is an important step in the understanding of processes involving coordinated cell behavior such as tissue development, wound healing, and cancer progression. Apart from extremely challenging in vivo studies, artificial tissue models are proven to be very useful in determining the main physical factors that affect the cooperativity of cells, simply because the conditions of growth can be very well controlled. One of the most established cell types in this field of research is the Madin-Darby canine kidney epithelial cell (MDCK), originating from the kidney distal tube (1). A great advantage of this polarized epithelial cell line is that it retained the ability for contact inhibition (2), which makes it a perfect model system for studies of epithelial morphogenesis.Organization of MDCK cells in colonies have been studied in a number of circumstances. For example, it was shown that in three-dimensional soft Matrigel, MDCK cells form a spherical enclosure of a lumen that is enfolded by one layer of polarized cells with an apical membrane exposed to the lumen side (3). These structures can be altered by introducing the hepatocyte growth factor, which induces the formation of linear tubes (4). However, the best-studied regime of growth is performed on two-dimensional surfaces where MDCK II cells form sheets and exhibit contact inhibition. Consequently, the obtained monolayers are well characterized in context of development (5), mechanical properties (6), and obstructed cell migration (7–9).Surprisingly, in the context of mechanics, several studies of monolayer formation showed that different rigidities of polydimethylsiloxane gels (5) and polyacrylamide (PA) gels (9) do not influence the nature of monolayer formation nor the attainable steady-state density. This is supposedly due to long-range forces between cells transmitted by the underlying elastic substrate (9). These results were found to agree well with earlier works on bovine aortic endothelial cells (10) and vascular smooth muscle cells (11), both reporting a lack of sensitivity of monolayers to substrate elasticity. Yet, these results are in stark contrast with single-cell experiments (12–15) that show a clear response of cell morphology, focal adhesions, and cytoskeleton organization to substrate elasticity. Furthermore, sensitivity to the presence of growth factors that are dependent on the elasticity of the substrate in two (16) and three dimensions (4) makes this result even more astonishing. Therefore, we readdress the issue of sensitivity of tissues to the elasticity of the underlying substrate and show that sufficiently soft gels induce a clearly different tissue organization.We plated MDCK II cells on soft PA gels (Young’s modulus E = 0.6 ± 0.2 kPa), harder PA gels (E = 5, 11, 20, 34 kPa), and glass, all coated with Collagen-I. Gels were prepared following the procedure described in Rehfeldt et al. (17); rigidity and homogeneity of the gels was confirmed by bulk and microrheology (see the Supporting Material for comparison). Seeding of MDCK II cells involved a highly concentrated solution dropped in the middle of a hydrated gel or glass sample. For single-cell experiments, cells were dispersed over the entire dish. Samples were periodically fixed up to Day 12, stained for nuclei and actin, and imaged with an epifluorescence microscope. Details are described in the Supporting Material.On hard substrates and glass it was found previously that the area of small clusters expands exponentially until the movement of the edge cannot keep up with the proliferation in the bulk (5). Consequently, the bulk density increases toward the steady state, whereas the density of the edge remains low. At the same time, the colony size grows subexponentially (5). This is what we denote “the classical regime of growth”. Our experiments support these observations for substrates with E ≥ 5 kPa. Specifically, on glass, colonies start as small clusters of very low density of 700 ± 200 cells/mm² (Fig. 1, A and B), typically surrounded by a strong actin cable (Fig. 1, B and C). Interestingly, the spreading area of single cells (Fig. 1 A) on glass was found to be significantly larger, i.e., (2.0 ± 0.9) × 10⁻³ mm². After Day 4 (corresponding cluster area of 600 ± 100 mm²), the density in the center of the colony reached the steady state with 6,800 ± 500 cells/mm², whereas the mean density of the edge profile grew to 4,000 ± 500 cells/mm². This density was retained until Day 12 (cluster area 1800 ± 100 mm²), which is in agreement with previous work (9).Open in a separate windowFigure 1Early phase of cluster growth on hard substrates. (A) Well-spread single cells, and small clusters with a visible actin cable 6 h after seeding. (B) Within one day, clusters densify and merge, making small colonies. (C) Edge of clusters from panel B.In colonies grown on 0.6 kPa gels, however, we encounter a very different growth scenario. The average spreading area of single cells is (0.34 ± 0.3) × 10⁻³ mm², which is six times smaller than on glass substrates (Fig. 2 A). Clusters of only few cells show that cells have a preference for cell-cell contacts (a well-established flat contact zone can be seen at the cell-cell interface in Fig. 2 A) rather than for cell-substrate contacts (contact zone is diffusive and the shape of the cells appears curved). The same conclusion emerges from the fact that dropletlike agglomerates, resting on the substrate, form spontaneously (Fig. 2 A), and that attempts to seed one single cluster of 90,000 cells fail, resulting in a number of three-dimensional colonies (Fig. 2 A). When the contact area with the substrate exceeds 4.7 × 10⁻³ mm², a monolayer appears in the center of such colonies (Fig. 2 B). The colonies can merge, and if individual colonies are small, the collapse into a single domain is associated with the formation of transient irregular structures (Fig. 2 B). Ultimately, large elliptical colonies (average major/minor axis of e = 1.8 ± 0.6) with a smooth edge are formed (Fig. 2 C), unlike on hard substrates where circular clusters (e = 1.06 ± 0.06) with a ragged edge comprise the characteristic phenotype.Open in a separate windowFigure 2Early phase of cluster growth on soft substrates. (A) Twelve hours after seeding, single cells remain mostly round and small. They are found as individual, or within small, three-dimensional structures (top). The latter nucleate a monolayer in their center (bottom), if the contact area with the substrate exceeds ∼5 × 10⁻³ mm². (B) Irregularly-shaped clusters appear due to merging of smaller droplets. A stable monolayer surrounded by a three-dimensional belt of densely packed cells is clearly visible, even in larger structures. (C) All colonies are recorded on Day 4.Irrespective of cluster size, in the new regime of growth, the internal structure is built of two compartments (Fig. 2 B):

• 1.The first is the edge (0.019 ± 0.05-mm wide), a three-dimensional structure of densely packed cells. This belt is a signature of the new regime because on hard substrates the edge is strictly two-dimensional (Fig. 1 C).
• 2.The other is the centrally placed monolayer with a spatially constant density that is very weakly dependent on cluster size and age (Fig. 3). The mean monolayer density is 13,000 ± 2,000 cells/mm², which is an average over 130 clusters that are up to 12 days old and have a size in the range of 10⁻³ to 10 mm², each shown by a data point in Fig. 3. This density is twice the steady-state density of the bulk tissue in the classical regime of growth.Open in a separate windowFigure 3Monolayer densities in colonies grown on 0.6 kPa substrates, as a function of the cluster size and age. Each cluster is represented by a single data point signifying its mean monolayer density. (Black lines) Bulk and (red dashed lines) edge of steady-state densities from monolayers grown on glass substrates. Error bars are omitted for clarity, but are discussed in the Supporting Material.

Until Day 4, the monolayer is very homogeneous, showing a nearly hexagonal arrangement of cells. From Day 4, however, defects start to appear in the form of small holes (typical size of (0.3 ± 0.1) × 10⁻³ mm²). These could be attributed to the extrusions of viable cells, from either the belt or areas of increased local density in the monolayer (inset in Fig. 4). This suggests that extrusions serve to release stress built in the tissue, and, as a consequence, the overall density is decreased.Open in a separate windowFigure 4Cell nuclei within the mother colony and in the neighboring archipelago of second-generation clusters grown on 0.6 kPa gels at Day 12. (Inset; scale bar = 10 μm) Scar in the tissue, a result of a cell-extrusion event. (Main image; scale bar = 100 μm) From the image of cell nuclei (left), it is clear that there are no cells within the scar, whereas the image of actin (right) shows that the cytoplasm of the cells at the edge has closed the hole.Previous reports suggest that isolated MDCK cells undergo anoikis 8 h after losing contact with their neighbors (18). However, in this case, it appears that instead of dying, the extruded cells create new colonies, which can be seen as an archipelago surrounding the mother cluster (Fig. 4). The viability of off-cast cells is further evidenced by the appearance of single cells and second-generation colonies with sizes varying over five orders of magnitude, from Day 4 until the end of the experiment, Day 12. Importantly, no morphological differences were found in the first- and second-generation colonies.In conclusion, we show what we believe to be a novel phase of growth of MDCK model tissue on soft PA gels (E = 0.6 kPa) that, to our knowledge, despite previous similar efforts (9), has not been observed before. This finding is especially interesting in the context of elasticity of real kidneys, for which a Young’s modulus has been found to be between 0.05 and 5 kPa (19,20). This coincides with the elasticity of substrates studied herein, and opens the possibility that the newly found phase of growth has a particular biological relevance. Likewise, the ability to extrude viable cells may point to a new migratory pathway regulated mechanically by the stresses in the tissue, the implication of which we hope to investigate in the future.     

3D-SIM Super-resolution of FtsZ and Its Membrane Tethers in Escherichia coli Cells

FtsZ, a bacterial homolog of eukaryotic tubulin, assembles into the Z ring required for cytokinesis. In Escherichia coli, FtsZ interacts directly with FtsA and ZipA, which tether the Z ring to the membrane. We used three-dimensional structured illumination microscopy to compare the localization patterns of FtsZ, FtsA, and ZipA at high resolution in Escherichia coli cells. We found that FtsZ localizes in patches within a ring structure, similar to the pattern observed in other species, and discovered that FtsA and ZipA mostly colocalize in similar patches. Finally, we observed similar punctate and short polymeric structures of FtsZ distributed throughout the cell after Z rings were disassembled, either as a consequence of normal cytokinesis or upon induction of an endogenous cell division inhibitor.The assembly of the bacterial tubulin FtsZ has been well studied in vitro, but the fine structure of the cytokinetic Z ring it forms in vivo is not well defined. Super-resolution microscopy methods including photoactivated localization microscopy (PALM) and three-dimensional-structured illumination microscopy (3D-SIM) have recently provided a more detailed view of Z-ring structures. Two-dimensional PALM showed that Z rings in Escherichia coli are likely composed of loosely-bundled dynamic protofilaments (1,2). Three-dimensional PALM studies of Caulobacter crescentus initially showed that Z rings were comprised of loosely bundled protofilaments forming a continuous but dynamic ring (1–3). However, a more recent high-throughput study showed that the Z rings of this bacterium are patchy or discontinuous (4), similar to Z rings of Bacillus subtilis and Staphylococcus aureus using 3D-SIM (5). Strauss et al. (5) also demonstrated that the patches in B. subtilis Z rings are highly dynamic.Assembly of the Z ring is modulated by several proteins that interact directly with FtsZ and enhance assembly or disassembly (6). For example, FtsA and ZipA promote ring assembly in E. coli by tethering it to the cytoplasmic membrane (7,8). SulA is an inhibitor of FtsZ assembly, induced only after DNA damage, which sequesters monomers of FtsZ to prevent its assembly into a Z ring (9). Our initial goals were to visualize Z rings in E. coli using 3D-SIM, and then examine whether any FtsZ polymeric structures remain after SulA induction. We also asked whether FtsA and ZipA localized in patchy patterns similar to those of FtsZ.We used a DeltaVision OMX V4 Blaze microscope (Applied Precision, GE Healthcare, Issaquah, WA) to view the high-resolution localization patterns of FtsZ in E. coli cells producing FtsZ-GFP (Fig. 1). Three-dimensional views were reconstructed using softWoRx software (Applied Precision). To rule out GFP artifacts, we also visualized native FtsZ from a wild-type strain (WM1074) by immunofluorescence (IF).Open in a separate windowFigure 1Localization of FtsZ in E. coli. (A) Cell with a Z ring labeled with FtsZ-GFP. (B) Rotated view of Z ring in panel A. (C) Cell with a Z ring labeled with DyLight 550 (Thermo Fisher Scientific, Waltham, MA). (D) Rotated view of Z ring in panel C. (B1 and D1) Three-dimensional surface intensity plots of Z rings in panels B and D, respectively. (E) A dividing cell producing FtsZ-GFP. The cell outline is shown in the schematic. (Asterisk) Focus of FtsZ localization; (open dashed ovals) filamentous structures of FtsZ. Three-dimensional surface intensity plots were created using the software ImageJ (19). Scale bars, 1 μm.Both FtsZ-GFP (Fig. 1, A, B, and B1) and IF staining for FtsZ (Fig. 1, C, D, and D1) consistently localized to patches around the ring circumference, similar to the B. subtilis and C. crescentus FtsZ patterns (4,5). Analysis of fluorescence intensities (see Fig. S1, A and B, in the Supporting Material) revealed that the majority of Z rings contain one or more gaps in which intensity decreases to background levels (82% for FtsZ-GFP and 69% for IF). Most rings had 3–5 areas of lower intensity, but only a small percentage of these areas had fluorescence below background intensity (34% for FtsZ-GFP and 21% for IF), indicating that the majority of areas with lower intensity contain at least some FtsZ.To elucidate how FtsZ transitions from a disassembled ring to a new ring, we imaged a few dividing daughter cells before they were able to form new Z rings (Fig. 1 E). Previous conventional microscopy had revealed dynamic FtsZ helical structures (10), but the resolution had been insufficient to see further details. Here, FtsZ visualized in dividing cells by 3D-SIM localized throughout as a mixture of patches and randomly-oriented short filaments (asterisk and dashed oval in Fig. 1, respectively). These structures may represent oligomeric precursors of Z ring assembly.To visualize FtsZ after Z-ring disassembly another way, we overproduced SulA, a protein that blocks FtsZ assembly. We examined E. coli cells producing FtsZ-GFP after induction of sulA expression from a pBAD33-sulA plasmid (pWM1736) with 0.2% arabinose. After 30 min of sulA induction, Z rings remained intact in most cells (Fig. 2 A and data not shown). The proportion of cellular FtsZ-GFP in the ring before and after induction of sulA was consistent with previous data (data not shown) (1,11).Open in a separate windowFigure 2Localization of FtsZ after overproduction of SulA. (A) Cell producing FtsZ-GFP after 0.2% arabinose induction of SulA for 30 min. (B) After 45 min. (B1) Magnified cell shown in panel B. (C) Cell producing native FtsZ labeled with AlexaFluor 488 (Life Technologies, Carlsbad, CA) 30 min after induction; (D) 45 min after induction. (D1) Magnified cell shown in panel D. Scale bars, 1 μm. (Asterisk) Focus of FtsZ localization; (open dashed ovals) filamentous structures of FtsZ.Notably, after 45 min of sulA induction, Z rings were gone (Fig. 2, B and B1), replaced by numerous patches and randomly-oriented short filaments (asterisk and dashed ovals in Fig. 2), similar to those observed in a dividing cell. FtsZ normally rapidly recycles from free monomers to ring-bound polymers (11), but a critical concentration of SulA reduces the pool of available FtsZ monomers, resulting in breakdown of the Z ring (9). The observed FtsZ-GFP patches and filaments are likely FtsZ polymers that disassemble before they can organize into a ring.We confirmed this result by overproducing SulA in wild-type cells and detecting FtsZ localization by IF (Fig. 2, C, D, and D1). The overall fluorescence patterns in cells producing FtsZ-GFP versus cells producing only native FtsZ were similar (Fig. 2, B1 and D1), although we observed fewer filaments with IF, perhaps because FtsZ-GFP confers slight resistance to SulA, or because the increased amount of FtsZ in FtsZ-GFP producing cells might titrate the SulA more effectively.Additionally, we wanted to observe the localization patterns of the membrane tethers FtsA and ZipA. Inasmuch as both proteins bind to the same C-terminal conserved tail of FtsZ (12–14), they would be expected to colocalize with the circumferential FtsZ patches in the Z ring. We visualized FtsA using protein fusions to mCherry and GFP (data not shown) as well as IF using a wild-type strain (WM1074) (Fig. 3 A). We found that the patchy ring pattern of FtsA localization was similar to the FtsZ pattern. ZipA also displayed a similar patchy localization in WM1074 by IF (Fig. 3 B).Open in a separate windowFigure 3Localization of FtsA (A) and ZipA (B) by IF using AlexaFluor 488. (C) FtsA-GFP ring. (D) Same cell shown in panel C with ZipA labeled with DyLight 550. (C1 and D1) Three-dimensional surface intensity plots of FtsA ring from panel C or ZipA ring from panel D, respectively. (E) Merged image of FtsA (green) and ZipA (red) from the ring shown in panels C and D. (F) Intensity plot of FtsA (green) and ZipA (red) of ring shown in panel E. The plot represents intensity across a line drawn counterclockwise from the top of the ring around the circumference, then into its lumen. Red/green intensity plot and three-dimensional surface intensity plots were created using the software ImageJ (19). Scale bar, 1 μm.To determine whether FtsA and ZipA colocalized to these patches, we used a strain producing FtsA-GFP (WM4679) for IF staining of ZipA using a red secondary antibody. FtsA-GFP (Fig. 3 C) and ZipA (Fig. 3 D) had similar patterns of fluorescence, although the three-dimensional intensity profiles (Fig. 3, C1 and D1) reveal slight differences in intensity that are also visible in a merged image (Fig. 3 E). Quantitation of fluorescence intensities around the circumference of the rings revealed that FtsA and ZipA colocalized almost completely in approximately half of the rings analyzed (Fig. 3 F, and see Fig. S2 A), whereas in the other rings there were significant differences in localization in one or more areas (see Fig. S2 B). FtsA and ZipA bind to the same C-terminal peptide of FtsZ and may compete for binding. Cooperative self-assembly of FtsA or ZipA might result in large-scale differential localization visible by 3D-SIM.In conclusion, our 3D-SIM analysis shows that the patchy localization of FtsZ is conserved in E. coli and suggests that it may be widespread among bacteria. After disassembly of the Z ring either in dividing cells or by excess levels of the cell division inhibitor SulA, FtsZ persisted as patches and short filamentous structures. This is consistent with a highly dynamic population of FtsZ monomers and oligomers outside the ring, originally observed as mobile helices in E. coli by conventional fluorescence microscopy (10) and by photoactivation single-molecule tracking (15). FtsA and ZipA, which bind to the same segment of FtsZ and tether it to the cytoplasmic membrane, usually display a similar localization pattern to FtsZ and each other, although in addition to the differences we detect by 3D-SIM, there are also likely differences that are beyond its ∼100-nm resolution limit in the X,Y plane.As proposed previously (16), gaps between FtsZ patches may be needed to accommodate a switch from a sparse Z ring to a more condensed ring, which would provide force to drive ring constriction (17). If this model is correct, the gaps should close upon ring constriction, although this may be beyond the resolution of 3D-SIM in constricted rings. Another role for patches could be to force molecular crowding of low-abundance septum synthesis proteins such as FtsI, which depend on FtsZ/FtsA/ZipA for their recruitment, into a few mobile supercomplexes.How are FtsZ polymers organized within the Z-ring patches? Recent polarized fluorescence data suggest that FtsZ polymers are oriented both axially and circumferentially within the Z ring in E. coli (18). The seemingly random orientation of the non-ring FtsZ polymeric structures we observe here supports the idea that there is no strong constraint requiring FtsZ oligomers to follow a circumferential path around the cell cylinder. The patches of FtsZ in the unperturbed E. coli Z ring likely represent randomly oriented clusters of FtsZ filaments that are associated with ZipA, FtsA, and essential septum synthesis proteins. New super-resolution microscopy methods should continue to shed light on the in vivo organization of these protein assemblies.     

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.