A +TIP for a smooth trip

Is there a cellular mechanism for preventing a depolymerizing microtubule track from “slipping out from under” its cargo? A recent study in budding yeast indicates that when a chromosome is transported to the minus end of a spindle microtubule, its kinetochore-bound microtubule plus end–tracking protein (+TIP) Stu2 may move to the plus end to promote rescue; i.e., to switch the depolymerizing end to a polymerizing end. The possibility that other +TIPs may play a similar role in sustaining a microtubule track during vesicular transport deserves investigation.Microtubule motor proteins such as dynein and kinesins are responsible for transporting cellular cargos along microtubule tracks (Vale, 2003). The net direction and speed of cargo movement, however, are likely to be regulated in a very complicated fashion, especially when a cargo is bound to multiple motors with opposite directionalities (Vale 2003; Mallik and Gross 2004; Levi et al., 2006). The fact that the microtubule track is not very stable further complicates matters. The plus ends of microtubules, which face the cell periphery in most cell types, are highly dynamic, exhibiting alternating periods of polymerization (growth) and depolymerization (shrinkage; Desai and Mitchison, 1997). Such plus end dynamics may be useful for searching and capturing relatively stationary cargos near the cell periphery that need to be transported inward (Vaughan et al., 2002). However, the dynamic nature of the track can also create an obvious problem for the transport process. If a microtubule''s rate of shrinkage is greater than the rate of cargo transport, then the microtubule may shrink past an attached cargo, causing its dissociation from the track. Does this happen in cells, or do cells have a mechanism to prevent it?Although this question has never been directly addressed, a recent study on budding yeast chromosome segregation has shed new light on the issue (Tanaka et al., 2005). In this study, the authors took advantage of a strategy that allowed them to specifically shut off the function of a single kinetochore, thereby preventing it from attaching to a spindle microtubule while, at the same time, permitting other kinetochores to attach to the spindle. After the function of this single kinetochore was switched back on, the behavior of its associated chromosome on a spindle microtubule was subjected to a detailed image analysis. Several important insights from this study on chromosome–microtubule interactions during mitosis have been recently reviewed (Bloom 2005), and, thus, only those observations that pertain to cargo transport will be highlighted here. The chromosome was first seen to undergo a lateral interaction with the microtubule followed by minus end–directed transport toward the pole. The mechanism of the minus end–directed transport is not entirely clear, although a member of the kinesin-14 family, Kar3, may be one of the players in this process (Tanaka et al., 2005). During transport, the attached microtubule can undergo shrinkage with a rate higher than that of the minus end–directed chromosome movement (Tanaka et al., 2005); however, it never shrank beyond the position of the cargo. Such exquisite control over the extent of shrinkage appears to rely on a conversation between the cargo and the plus end of the microtubule that is mediated by the microtubule plus end–tracking protein Stu2 (Bloom 2005; Tanaka et al., 2005).Microtubule plus end–tracking proteins (+TIPs) are a class of proteins that use different structural motifs or specific targeting mechanisms to localize to the dynamic plus ends of microtubules (Carvalho et al., 2003; Akhmanova and Hoogenraad, 2005). Although most +TIPs associate with only the growing ends of microtubules, several +TIPs also localize to the shrinking ends (Carvalho et al., 2003; 2004; Akhmanova and Hoogenraad, 2005; Mennella et al., 2005; Sproul et al., 2005; Molk et al., 2006; Wu et al., 2006). Many +TIPs have been found to impact microtubules by either promoting their growth or promoting dynamic behavior. Stu2 is a member of the XMAP215/TOG/Dis1/DdCP224 family of proteins that have been shown to affect microtubule dynamics in multiple ways depending on different experimental conditions (Ohkura et al., 2001; Popov and Karsenti 2003; Holmfeldt et al., 2004; Akhmanova and Hoogenraad, 2005). In vitro, Stu2 binds to the plus ends of preformed microtubules and promotes catastrophe, which is a switch from growth to shrinkage (van Breugel et al., 2003). In vivo studies using mutants of Stu2, however, indicate that Stu2 promotes microtubule growth (Severin et al., 2001) and the dynamics of both kinetochore and cytoplasmic microtubules (Kosco et al., 2001; Pearson et al., 2003). During anaphase B spindle elongation, Stu2 may antagonize the function of Kip3 (a kinesin-13 family member) to promote the plus end polymerization of overlapping microtubules (Severin et al., 2001). Although it is not fully understood how or why Stu2 is so versatile, it is well recognized that the in vivo interactions among +TIPs are very complicated, and the loss of function of a +TIP in vivo may decrease or increase the accumulation of other +TIPs that also regulate microtubule dynamics (Carvalho et al., 2003; 2004; Lansbergen et al., 2004; Akhmanova and Hoogenraad, 2005; Galjart 2005; Komarova et al., 2005). Tanaka et al. (2005) identified Stu2 as a rescue (a switch from shrinkage to growth) factor based not on phenotypic studies of Stu2 mutants but, instead, on a direct observation of the relationship between microtubule plus end behavior and Stu2 localization. They found that Stu2 was localized at the plus ends of microtubules emanating from the spindle pole body, and, during periods of microtubule shrinkage, Stu2 levels at the plus ends were decreased. Interestingly, Stu2 was also localized at the unbound kinetochore. When the kinetochore subsequently attached laterally to a spindle microtubule and underwent minus end–directed transport, the Stu2 proteins were transported from the kinetochore to the microtubule plus end. The arrival of Stu2 at the plus end closely correlated to the rescue of the shrinking microtubule (Tanaka et al., 2005). These observations strongly suggest that the Stu2 carried by the kinetochore may serve as a rescue factor for the microtubule track, preventing it from vanishing before the migrating chromosome.Could such a scenario exist during microtubule-dependent transport of nonchromosomal cargoes during interphase? We do not yet know the answer. However, based on published studies, it seems reasonable to hypothesize that other +TIPs, especially the cytoplasmic linker protein CLIP-170, may function in a manner similar to yeast Stu2 to ensure a safe trip for a minus end–directed cargo. CLIP-170 contains CAP-Gly microtubule-binding motifs at its NH₂ terminus and was initially identified as a protein required for linking endocytic vesicles to microtubules in vitro (Pierre et al., 1992; Rickard and Kreis 1996). Later, CLIP-170 was identified as a founding member of the microtubule plus end–tracking proteins (Perez et al., 1999). The connection between CLIP-170''s in vitro endosome–microtubule linking property and its in vivo plus end tracking behavior has not been clearly made. Could an endocytic vesicle use its bound CLIP-170 as a rescue factor to prevent the disappearance of the track on which it is traveling?CLIP-170 is indeed considered to be a rescue factor in mammalian cells (Komarova et al., 2002a). Komarova et al. (2002b) have found that in cultured CHO and NRK cells, microtubule dynamics seem to be controlled spatially; catastrophe and rescue occur frequently only near the cell periphery. Although the mechanisms behind catastrophe and rescue are not fully understood, protein factors are required for regulating both events in vivo (Desai and Mitchison, 1997). In CHO cells, a dominant-negative form of CLIP-170 that displaces the endogenous CLIP-170 from microtubule plus ends severely reduces the rescue frequency so that microtubules are more likely to shrink all the way back to the microtubule-organizing center (Komarova et al., 2002a). Moreover, both in vivo and in vitro studies suggest that the rescue activity of CLIP-170 is localized to the NH₂ terminus containing the CAP-Gly motifs (Komarova et al., 2002a; Arnal et al., 2004). How CLIP-170 rescues a shrinking end is not clear. CLIP-170 can promote tubulin oligomerization (Diamantopoulos et al., 1999; Arnal et al., 2004), and it is likely that this property serves to increase the local concentration of tubulin substrate, thereby lowering the entropic barrier for the polymerization reaction. CLIP-170 in mammalian cells has only been found at growing plus ends, most likely as a result of copolymerization with tubulin subunits followed by its release from older segments (Diamantopoulos et al., 1999; Perez et al., 1999; Folker et al., 2005). When a microtubule end shrinks, CLIP-170 falls off. Is there a mechanism to get CLIP-170 close to the depolymerizing end and facilitate its function as a rescue factor? Given the proposed function of Stu2 as a rescue factor for spindle microtubules, one may easily imagine a similar scenario in which vesicle-bound CLIP-170 may be transported to the approaching microtubule end to rescue it from further shrinkage.If vesicle-bound CLIP-170 is transported to the plus end in a manner similar to Stu2, could such transport be mediated by plus end–directed kinesins? Although the kinesin involved in transporting Stu2 toward the microtubule plus end still needs to be identified, detailed image analyses have revealed a role for the Kip2/Tea2 kinesins (members of the kinesin-7 family) in transporting CLIP-170 homologues in fungi (Busch et al., 2004; Carvalho et al., 2004). Bik1 and Tip1 are the CLIP-170 homologues in budding and fission yeasts, respectively, and these proteins are found at microtubule plus ends, where they act as growth-promoting factors or anticatastrophe factors (Berlin et al., 1990; Brunner and Nurse 2000; Carvalho et al., 2004). In both yeasts, the Kip2/Tea2 kinesins bind to and comigrate with the CLIP-170 homologues along the microtubule toward the plus end (Busch et al., 2004; Carvalho et al., 2004). Kinesins have also been implicated in targeting other +TIPs to microtubule plus ends (Jimbo et al., 2002; Maekawa et al., 2003; Zhang et al., 2003; Wu et al., 2006). For example, the mammalian tumor suppressor protein APC (adenomatous polyposis coli) may be targeted to the plus end by KIF3A/KIF3B (a heterotrimeric kinesin II in the kinesin-2 family) as well as by other mechanisms (Jimbo et al., 2002; Nathke 2004; Slep et al., 2005). It will be interesting to see whether a similar transport process for CLIP-170 exists in higher eukaryotic cells. It is possible that such a mechanism would deliver just enough CLIP-170 to the shrinking plus end to initiate rescue. When microtubule growth is resumed, CLIP-170''s intrinsic higher affinity for tubulin subunits and lower affinity for the microtubule wall may allow these proteins to “treadmill” on the growing end (Perez et al., 1999; Folker et al., 2005).The regulation of CLIP-170 activity appears to be rather complex. CLIP-170 is most likely phosphorylated by multiple kinases, including FKBP12–rapamycin-associated protein (mTOR; Choi et al., 2002). Although phosphorylation by mTOR/FKBP 12–rapamycin-associated protein may stimulate CLIP-170''s microtubule binding, phosphorylation by other kinases may cause CLIP-170 to dissociate from microtubules (Rickard and Kreis 1996; Choi et al., 2002). In vivo, CLIP-170 has a closed conformation that is presumably inactive and an open conformation that may interact with microtubules and dynein regulators such as dynactin (Schroer 2004) and LIS1 (Morris et al., 1998; Lansbergen et al., 2004). It is possible that phosphorylation may regulate the conversion between these two forms, but the specific mechanism and the spatial regulation for this conversion have yet to be resolved. If CLIP-170 is indeed released from a membranous cargo to move to the plus end in order to serve as a rescue factor, it would be interesting to know when and/or where such a conformational switch occurs. Finally, other proteins may play redundant roles with CLIP-170 in vesicular trafficking, which may explain why a dramatic defect in vesicle/organelle distribution is not detected when the CLIP-170 level is lowered or when the gene is knocked out (Lansbergen et al., 2004; Akhmanova et al., 2005).+TIPs other than CLIP-170 may play a similar role in rescuing shrinking microtubule tracks. For example, the dynactin complex that links dynein to membranous cargoes and promotes the processive motion of dynein (Schroer 2004) may act as a rescue factor. The p150^Glued subunit of dynactin and CLIP-170 both contain CAP-Gly microtubule-binding motifs at their NH2 termini, although p150^Glued contains one, whereas CLIP-170 contains two such motifs. Dynactin has been shown to behave as a +TIP facilitating the capture of vesicular cargo for minus end–directed transport (Vaughan et al., 1999; 2002). The head domain of the p150^Glued subunit containing the CAP-Gly motif has been shown to promote rescue in vivo in the absence of endogenous CLIP-170, although the effect was much weaker than that caused by the exogenous CLIP-170 head domain (Kamarova et al., 2002a). In vitro studies showed that dynactin may promote nucleation during microtubule assembly (Ligon et al., 2003), which is consistent with it being a potential rescue factor. As shown with CLIP-170, this capacity to bring multiple tubulins together may help to overcome the entropic barrier of the polymerization reaction. Finally, cargo-bound dynactin may also use kinesin to get to the plus end. The p150^Glued subunit of dynactin has been shown to interact directly with the COOH terminus of KAP3, a subunit of the heterotrimeric kinesin II (a member of the kinesin-2 family) that also binds to APC (Jimbo et al., 2002; Deacon et al., 2003; Dell 2003). Although this binding is implicated in dynactin''s role as a cargo adaptor for kinesin II, it is possible, in theory, that a small amount of dynactin may use this connection to move to the plus end.The proposed hypothesis that +TIPs may be released from a membranous cargo to rescue a shrinking microtubule track may apply to both minus and plus end–directed transport. In addition, it is important to point out that this hypothesis does not exclude other mechanisms for rescuing long microtubule tracks. Rescue may occur stochastically, and, sometimes, +TIPs may participate in other ways such as mediating microtubule capture by the actin-rich cortex to stabilize the track (Wen et al., 2004; Galjart 2005). In some situations, microtubule dynamics are modulated by the direct binding of membranous cargo to the growing or shrinking plus ends of microtubules (Waterman-Storer and Salmon, 1998).Currently, the ability of CLIP-170 or other +TIPs to be released from a membranous cargo and to act as a rescue factor for a shrinking microtubule is just a hypothesis. Nevertheless, searching for proteins involved in the communication between a cargo and the approaching shrinking end of its microtubule track is clearly an endeavor worth pursuing.     

Cortical Microtubule Arrays Are Initiated from a Nonrandom Prepattern Driven by Atypical Microtubule Initiation

The ordered arrangement of cortical microtubules in growing plant cells is essential for anisotropic cell expansion and, hence, for plant morphogenesis. These arrays are dismantled when the microtubule cytoskeleton is rearranged during mitosis and reassembled following completion of cytokinesis. The reassembly of the cortical array has often been considered as initiating from a state of randomness, from which order arises at least partly through self-organizing mechanisms. However, some studies have shown evidence for ordering at early stages of array assembly. To investigate how cortical arrays are initiated in higher plant cells, we performed live-cell imaging studies of cortical array assembly in tobacco (Nicotiana tabacum) Bright Yellow-2 cells after cytokinesis and drug-induced disassembly. We found that cortical arrays in both cases did not initiate randomly but with a significant overrepresentation of microtubules at diagonal angles with respect to the cell axis, which coincides with the predominant orientation of the microtubules before their disappearance from the cell cortex in preprophase. In Arabidopsis (Arabidopsis thaliana) root cells, recovery from drug-induced disassembly was also nonrandom and correlated with the organization of the previous array, although no diagonal bias was observed in these cells. Surprisingly, during initiation, only about one-half of the new microtubules were nucleated from locations marked by green fluorescent protein-γ-tubulin complex protein2-tagged γ-nucleation complexes (γ-tubulin ring complex), therefore indicating that a large proportion of early polymers was initiated by a noncanonical mechanism not involving γ-tubulin ring complex. Simulation studies indicate that the high rate of noncanonical initiation of new microtubules has the potential to accelerate the rate of array repopulation.Higher plant cells feature ordered arrays of microtubules at the cell cortex (Ledbetter and Porter, 1963) that are essential for cell and tissue morphogenesis, as revealed by disruption of cortical arrays by drugs that cause microtubule depolymerization (Green, 1962) or stabilization (Weerdenburg and Seagull, 1988) and by loss-of-function mutations in a wide variety of microtubule-associated proteins (Baskin, 2001; Whittington et al., 2001; Buschmann and Lloyd, 2008; Lucas et al., 2011). The structure of these arrays is thought to control the pattern of cell growth primarily by its role in the deposition of cellulose microfibrils, the load-bearing component of the cell wall (Somerville, 2006). Functional relations between cortical microtubules and cellulose microfibrils have been proposed since the early sixties, even before cortical microtubules had been visualized (Green, 1962). Recent live-cell imaging studies have confirmed that cortical microtubules indeed guide the movement of cellulose synthase complexes that produce cellulose microfibrils (Paredez et al., 2006) and have shown further that microtubules position the insertion of most cellulose synthase complexes into the plasma membrane (Gutierrez et al., 2009). These activities of ordered cortical microtubules are proposed to facilitate the organization of cell wall structure, creating material properties that underlie cell growth anisotropy.While organization of the interphase cortical array appears to be essential for cell morphogenesis, this organization is disrupted during the cell cycle as microtubules are rearranged to create the preprophase band, spindle, and phragmoplast during mitosis and cytokinesis (for review, see Wasteneys, 2002). Upon completion of cytokinesis, an organized interphase cortical array is regenerated, but the pathway for this reassembly is not well understood.The plant interphase microtubule array is organized and maintained without centrosomes as organizing centers (for review, see Wasteneys, 2002; Bartolini and Gundersen, 2006; Ehrhardt and Shaw, 2006), and microtubule self-organization is proposed to play an important role in cortical microtubule array ordering (Dixit and Cyr, 2004). In electron micrographs, microtubules have been observed to be closely associated with the plasma membrane (Hardham and Gunning, 1978), and live-cell imaging provides evidence for attachment of microtubules to the cell cortex (Shaw et al., 2003; Vos et al., 2004). The close association to the plasma membrane restricts the cortical microtubules to a quasi two-dimensional plane where they interact through polymerization-driven collisions (Shaw et al., 2003; Dixit and Cyr, 2004). Microtubule encounters at shallow angles (<40°) have a high probability of leading to bundling, while microtubule encounters at steeper angles most likely result in induced catastrophes or microtubule crossovers (Dixit and Cyr, 2004). Several computational modeling studies have since shown that these types of interactions between surface-bound dynamical microtubules can indeed explain spontaneous coalignment of microtubules (Allard et al., 2010; Eren et al., 2010; Hawkins et al., 2010; Tindemans et al., 2010).The question of how the orientation of the cortical array is established with respect to the cell axis is less well understood. One possibility is that microtubules are selectively destabilized with respect to cellular coordinates (Ehrhardt and Shaw, 2006). Indeed, recent results from biological observations and modeling suggest that catastrophic collisions induced at the edges between cell faces or heighted catastrophe rates in cell caps could be sufficient to selectively favor microtubules in certain orientation and hence determine the final orientation of the array (Allard et al., 2010; Eren et al., 2010; Ambrose et al., 2011; Dhonukshe et al., 2012).To date, all models of cortical array assembly assume random initial conditions. However, experimental work by Wasteneys and Williamson (1989a, 1989b) in Nitella tasmanica showed that, during array reassembly after drug-induced disruption, microtubules were initially transverse. This was followed by a less ordered phase and later by the acquisition of the final transverse order. A nonrandom initial ordering was also observed in tobacco (Nicotiana tabacum) Bright Yellow-2 (BY-2) cells by Kumagai et al. (2001), who concluded that the process of transverse array establishment starts with longitudinal order but did not provide quantitative data for the process of array assembly. The initial conditions for the cortical microtubule array formation are important to consider, as they may strongly influence the speed at which order is established and could even prevent it from being established over a biologically relevant time scale.In this study, we used live-cell imaging to follow and record the whole transition from the cortical microtubule-free state to the final transverse array and used digital tracking algorithms to quantify the microtubule order. Nucleation stands out as a central process to characterize during array initiation. Lacking a central body to organize microtubule nucleations, the higher plant cell has dispersed nucleation complexes (Wasteneys and Williamson, 1989a, 1989b; Chan et al., 2003; Shaw et al., 2003; Murata et al., 2005; Pastuglia et al., 2006; Nakamura et al., 2010). Therefore, we performed high time resolution observations to quantify nucleation complex recruitment, nucleation rates, and microtubule nucleation angles. We found evidence for a highly nonrandom initial ordering state that features diagonal microtubule orientation and an atypical microtubule initiation mechanism. Simulation analysis indicates that these atypical nucleations have the potential to accelerate the recovery of cortical array density.     

The functions and consequences of force at kinetochores

Chromosome segregation requires the generation of force at the kinetochore—the multiprotein structure that facilitates attachment of chromosomes to spindle microtubules. This force is required both to move chromosomes and to signal the formation of proper bioriented attachments. To understand the role of force in these processes, it is critical to define how force is generated at kinetochores, the contributions of this force to chromosome movement, and how the kinetochore is structured and organized to withstand and respond to force. Classical studies and recent work provide a framework to dissect the mechanisms, functions, and consequences of force at kinetochores.Force plays key roles in many different cellular processes by influencing objects in a way that causes them to change their speed or direction of movement. Force can take multiple forms in a cell and have very different consequences, depending on the circumstances of its action. When force pulls on an object, it creates “tension.” In contrast, a pushing force exerted upon an object is termed “compression.” To understand the contribution of force to cellular processes, it is important to determine the molecular mechanisms by which force is generated or produced at a subcellular structure, how these structures withstand the force, and how they detect and signal the presence of force. The process of mitotic chromosome segregation provides a particularly intriguing example of the importance of cellular force. During mitosis, force plays a critical role in directing the physical segregation of chromosomes and modulating the signals that sense and promote their proper attachment to the spindle. The central player in chromosome segregation is a macromolecular structure termed the kinetochore that establishes and maintains the attachment of each set of paired sister chromatids to microtubule polymers from opposing spindle poles and directs the segregation of chromosomes to the daughter cells (Cheeseman and Desai, 2008; Santaguida and Musacchio, 2009). The kinetochore plays key roles throughout mitosis, both to mediate direct attachments between microtubules and centromeric DNA (Fig. 1) and as a hub for the signaling molecules required to monitor and control faithful chromosome segregation and cell cycle progression. Because the kinetochore is the contact point between chromosomes and microtubules, the forces derived from microtubules are exerted directly on the proteins within the kinetochore. A key challenge is to understand how this force is generated and accommodated and to define the specific contributions of this force to kinetochore function.Open in a separate windowFigure 1.Simplified diagram of the kinetochore showing the major proteins involved in the DNA–microtubule attachment. (Left) The Ndc80 complex (dark blue) binds to microtubules and forms two separate connections to kinetochores. First, the Ndc80 complex binds to the Mis12 complex (green) and KNL-1 (magenta). The Mis12 complex in turn binds to CENP-C (orange), which binds to nucleosomes containing the histone H3 variant CENP-A (purple). Second, the Ndc80 complex binds to CENP-T (light blue). CENP-T interacts with DNA as a part of a heterotetrameric nucleosome-like CENP-T–W–S–X complex. In humans, the Ndc80 complex attachment to microtubules is enhanced by an interaction with the Ska1 complex (pink and blue; Schmidt et al., 2012). Additional components may form interactions between the two connective pathways (red). (Right) Upon microtubule depolymerization, the flexible protein components of the kinetochore may rearrange. For example, recent evidence has suggested that the N and C termini of CENP-T separate under tension (Suzuki et al., 2011) and that the subunits of the Mis12 complex redistribute (Wan et al., 2009).

How much force is generated at a kinetochore?

The nature of the forces involved in partitioning chromosomes has been an active area of research for more than 50 years. Edwin Taylor and Bruce Nicklas were among the first to consider the forces that resist chromosome movement. Separate theoretical analyses predicted that ∼0.1 pN would be required to move a chromosome at 1 µm/min when resisted only by viscous cytoplasmic drag (Nicklas, 1965; Taylor, 1965). Almost 20 years after publishing his theoretical work, Nicklas was able to test the force on a single chromosome during anaphase of meiosis I (Nicklas, 1983). Using a microneedle to measure the stall force on chromosomes in grasshopper spermatocytes, Nicklas found that 700 pN could act on a chromosome (Nicklas, 1983). He estimated that the kinetochores tested in these studies were bound by ∼15 microtubules (Nicklas, 1983), suggesting that each microtubule may be capable of generating up to ∼45 pN of force. In a later study, Nicklas determined that ∼50 pN of force was produced on a chromosome during prometaphase (Nicklas, 1988). This calculation was based on observations of chromosome congression and correlations with his previous work. By Nicklas’ own admission, the microneedle assays to measure the force exerted on anaphase chromosomes had a high associated error, and it is unknown whether forces in the hundreds of piconormals would ever be produced at a kinetochore in the absence of a perturbation. Regardless, no other work since has provided a more exact measurement, and 700 pN remains the standard reference value for the force that can act at a metazoan kinetochore.As Nicklas’ work suggested, it is likely that the force felt by kinetochores varies throughout the cell cycle and under different types of attachments (discussed later in this paper). In particular, the arrangement of paired sister chromatids attached to opposite spindle poles during metaphase would allow for the greatest tension to be applied to kinetochores. Recent work visualizing sister chromatid oscillations during metaphase has observed that at time points immediately before the switch from poleward to antipoleward motion, the poleward-moving kinetochore experiences the highest forces, at least as judged by changes in intra- and interkinetochore distances (Dumont et al., 2012; Wan et al., 2012). In addition, the antipoleward-moving kinetochore may experience passive forces (Inoué and Salmon, 1995; Maddox et al., 2003) that can also alter inter- and intrakinetochore stretch (Dumont et al., 2012; Wan et al., 2012). However, the magnitude of force during these directional switches and how this force is accommodated continues to be a subject of debate. As the higher order organization of kinetochores remains unknown, it is unclear how the forces from the multiple microtubule interactions at a single kinetochore are combined or what force is experienced by an individual protein within the kinetochore structure.

The mechanisms of force production

With the discovery of the potentially large forces produced at kinetochores (Nicklas, 1983), a major challenge has been to define the mechanisms by which this force is generated. Many initial studies focused on the contributions of the microtubule-based motors, dynein and kinesin, that were found to localize to kinetochores (Inoué and Salmon, 1995). The ability of these motors to transport cargoes along microtubules suggested that they might function similarly to move a chromosome within a cell. Individual kinesin and dynein motors have been shown to stall under ∼5–7 pN of opposing force, termed a load (Visscher et al., 1999; Gennerich et al., 2007), and the combined action of multiple motors could generate the forces that Nicklas observed. However, subsequent studies have found that chromosome movement can still largely occur in the absence of these motors in fungi (Cottingham et al., 1999; Grishchuk and McIntosh, 2006). In metazoans, motors, including the kinesin CENP-E and dynein, contribute to chromosome segregation (Sharp et al., 2000; Kapoor et al., 2006; Yang et al., 2007), although their relative importance remains unclear. An alternative hypothesis was that the microtubules themselves generated the force to move chromosomes (Inoué and Salmon, 1995). Several early studies provided evidence that microtubules could direct the movement of isolated chromosomes under conditions that would not permit motor protein function (Koshland et al., 1988; Coue et al., 1991; Hunt and McIntosh, 1998). This microtubule-derived movement could be caused by forces generated either at the kinetochore by microtubule depolymerization (Grishchuk et al., 2005) or at the spindle poles as a result of poleward flux and microtubule disassembly at the minus end (LaFountain et al., 2001, 2004; Chen and Zhang, 2004). In fact, subsequent work suggested that the stall forces measured by Nicklas were a result of minus end microtubule disassembly in equilibrium with the plus end microtubule polymerization caused by the application of tension via the microneedle (LaFountain et al., 2001, 2004; Chen and Zhang, 2004). Although it is now generally accepted that microtubules generate the primary forces responsible for chromosome movement, kinetochore-localized motors may generate some force, act as a “back-up” system when kinetochore capture by microtubules fails (Kapoor et al., 2006), generate tension via the production of the polar ejection forces (Mazumdar and Misteli, 2005), function to distribute force over additional linkages, and regulate microtubule dynamics (Bader and Vaughan, 2010; Al-Bassam and Chang, 2011). In addition to forces generated either directly or indirectly by the microtubules, a third model proposes that the chromosomes themselves may contribute to the segregation process because of entropic forces that act on the DNA (Jun and Wright, 2010; Finan et al., 2011). Although such forces would likely be very small, they may assist chromosome distribution, particularly in smaller cells.In support of a primary role for microtubules in generating force at kinetochores, microtubules have been shown to generate pulling force during their depolymerization in vitro (Grishchuk et al., 2005; Powers et al., 2009; Akiyoshi et al., 2010; Tien et al., 2010). During microtubule polymerization, GTP-bound tubulin dimers are added to the growing microtubule plus end (Desai and Mitchison, 1997). After these dimers are incorporated into the microtubule lattice, GTP is hydrolyzed. The resulting GDP-bound tubulin dimers associate with each other along an individual protofilament and between neighboring protofilaments within the microtubule lattice to maintain a straight microtubule (Nogales, 2000; Nogales and Wang, 2006). However, when a microtubule switches to depolymerization, a process termed catastrophe, GDP-bound dimers exposed at the microtubule end lose these stabilizing interactions, causing the protofilaments to peel backward. According to measurements and calculations by Grishchuk et al. (2005), the conformational change that occurs for an individual depolymerizing protofilament can generate a power stroke of ≤5 pN, suggesting that a depolymerizing microtubule composed of 13 protofilaments could generate as much as 65 pN of force. Importantly, to harness this force and ensure proper chromosome movement, it is critical to control microtubule polymerization and depolymerization at kinetochores. The formation of kinetochore–microtubule attachments as well as the resulting tension may directly modulate microtubule dynamics by slowing microtubule depolymerization and decreasing the rate of catastrophe (Franck et al., 2007; Akiyoshi et al., 2010; Umbreit et al., 2012). In addition, microtubule polymerization factors, such as the TOG (tumor overexpressed gene) domain proteins XMAP215 and CLASP, and depolymerases, such as kinesin-13 proteins, which are present both at the kinetochore and on the spindle, also modulate microtubule behavior (Bader and Vaughan, 2010; Al-Bassam and Chang, 2011).Although microtubule depolymerization has the capacity to generate force, a key question is how chromosome movement is coupled to microtubule depolymerization. Thus far, two models have dominated the literature to explain how kinetochores harness the force from microtubule depolymerization, although these models are not mutually exclusive. The first model, termed the “Hill sleeve” model or “biased diffusion” (Hill, 1985), postulates that the association of the kinetochore with a microtubule is formed by multiple weak interactions that can diffuse equally in either direction. However, because of a large free energy barrier that disfavors the loss of an interaction, this diffusion is biased toward the microtubule minus end as binding sites disappear from the plus end. The second model, termed the “forced walk” model (Molodtsov et al., 2005), proposes that the kinetochore is coupled to microtubules in such a way that, as the protofilaments peel backward during depolymerization, the coupling protein is pushed along the microtubule. The way in which the microtubule is connected to the kinetochore has important implications for understanding how the force manifests at the kinetochore and remains an important focus for future work.Recent studies have focused on how kinetochores and kinetochore proteins harness the energy from microtubule depolymerization. These studies have tested key players at the kinetochore–microtubule interface, such as the Ndc80, Dam1, and Ska1 complexes (McIntosh et al., 2008; Powers et al., 2009; Welburn et al., 2009; Lampert et al., 2010; Tien et al., 2010; Schmidt et al., 2012) for their abilities to track on depolymerizing microtubules, and have attempted to analyze the kinetochore as a whole using partial purifications of kinetochores from Saccharomyces cerevisiae (Akiyoshi et al., 2010). Although individual protein complexes and isolated yeast kinetochores are able to move with depolymerizing microtubules, studies performed using optical tweezers have found that the tested proteins and complexes are able to withstand less than 10 pN of pulling force before a rupture event is observed (Powers et al., 2009; Akiyoshi et al., 2010; Tien et al., 2010). This is in contrast to the theoretical maximum of 65 pN that a microtubule has been proposed to produce during depolymerization (Grishchuk et al., 2005). It is likely that in the context of a kinetochore assembled on a chromosome, the complex architecture of the kinetochore has the capacity to harness and withstand larger forces. Thus, the in vivo load-bearing properties of the kinetochore likely depend on a combination of the properties of both the individual protein components and the organization of the entire complex.

Signaling the biorientated state of chromosomes

During mitosis, it is critical that paired sister chromatids attach to opposite spindle poles. When this biorientation fails, this error must be detected and corrected, and a signal to delay cell cycle progression must be produced to prevent chromosome missegregation. Work performed by Li and Nicklas (1995) and Nicklas et al. (1995) demonstrated that the external application of force to a chromosome using a microneedle could overcome the checkpoint signal generated by an unattached kinetochore. This and other work have supported the model that the tension produced on bioriented sister kinetochores can alter the signaling state of the kinetochore. This tension results in two apparent physical alterations to mitotic chromosome structure: an increase in the distance between paired sister kinetochores and an increase in the distance between the inner and outer kinetochore regions of a single kinetochore. Under some conditions, this inter- and intrakinetochore stretch can be uncoupled (Maresca and Salmon, 2009), and recent research has focused on the importance of intrakinetochore stretch in modulating the signals that monitor attachment state. By measuring the relative spatial positions of the different kinetochore proteins, work from several groups has found that kinetochore structure is altered when chromosomes are bioriented relative to conditions of reduced tension (Maresca and Salmon, 2009; Uchida et al., 2009; Wan et al., 2009; Suzuki et al., 2011; Dumont et al., 2012). Biorientation results in the separation of inner kinetochore components (such as CENP-A and CENP-C) from outer kinetochore components (such as Ndc80 and Mis12) as well as changes in the spatial distribution of other proteins within the kinetochore and possibly conformational changes within the proteins themselves.Because the generation of tension is dependent on the presence of opposing forces, changes in kinetochore structure correlate with the successful bioriented arrangement of chromosomes on the metaphase plate. In contrast, when one sister kinetochore lacks an attachment to the spindle (monotelic), or if both kinetochores attach to the same pole (syntelic), it is not possible to generate similar opposing forces. However, even in these cases, some force may still be present because of the viscosity of the cytoplasm resisting chromosome movement (Nicklas, 1965; Taylor, 1965) or the action of chromokinesins that generate polar ejection forces (Mazumdar and Misteli, 2005). It remains unclear how force is exerted on a single kinetochore that simultaneously attaches to opposing spindle poles (merotelic) or how these incorrect attachments are resolved (Gregan et al., 2011; Matos and Maiato, 2011). The observed structural changes at kinetochores have been assumed to correlate with the presence of tension, but thus far, such studies have not made direct measurements of force or tension. Nevertheless, careful quantitative analysis of the dynamic changes in the distances between CENP-C and Hec1 or Cdc20 during sister chromatid oscillations has supported the model that changes in intrakinetochore distance are force dependent (Dumont et al., 2012). However, these structural alterations may also be the result of changes in the conformation, organization, or localization of proteins within the kinetochore.Ultimately, it is important to translate the mechanical signals produced by force at kinetochores into a chemical signal that regulates the activities of kinetochore proteins. A key player in correcting errors in microtubule attachment state is the Aurora B kinase. Substrates for Aurora B show tension-sensitive phosphorylation; they are highly phosphorylated in the absence of tension and become dephosphorylated upon biorientation (Liu et al., 2009; Welburn et al., 2010). The forces generated at kinetochores have been implicated in controlling Aurora B signaling by altering the spatial separation between the kinase and its substrates (Tanaka, 2002; Liu et al., 2009), although other models for tension-sensitive Aurora B phosphorylation have also been proposed (Sandall et al., 2006). The key substrates of Aurora B are located at the outer kinetochore and can be >100 nm away from the majority of Aurora B, which is localized at the inner centromere, depending on whether the sister kinetochores are under tension (Wan et al., 2009). Therefore, structural changes caused by opposing force at kinetochores separate the kinase and its substrates. The increased separation under tension makes Aurora B less likely to phosphorylate its now distant substrates (Liu et al., 2009; Welburn et al., 2010). One effect of Aurora B phosphorylation on outer kinetochore proteins is to reduce their microtubule binding affinity (Cheeseman et al., 2006; Welburn et al., 2010; Schmidt et al., 2012). Thus, it has been proposed that the presence of tension can ultimately stabilize microtubule attachments through changes in kinetochore conformation that cause a decrease in Aurora B phosphorylation, which in turn increases the microtubule binding activities of various kinetochore components.In addition to altering the signaling state of kinetochores, changes in force at kinetochores may also have a direct effect on microtubule binding. One recent study suggested that outer kinetochore proteins are force sensitive and show catch–slip properties (Akiyoshi et al., 2010), resulting in less frequent detachment under increasing force. This is analogous to a “Chinese finger trap” and would allow the attachment to become stabilized as the microtubule pulls on the kinetochore. Whether tension affects kinetochore–microtubule attachments directly or indirectly, force appears to play an essential role in establishing and signaling biorientation in addition to driving chromosome movement.

Theoretical considerations for force resistance

Force is a vector quantity that, when applied to a bond, decreases bond energy barriers, increasing the likelihood of bond breakage. Although the kinetochore must function under force to perform its roles properly, this force also represents a challenge with the potential for deleterious consequences to kinetochore function. Force could result in protein unfolding or the breakage of protein–protein interactions (Fig. 2). If a core kinetochore protein unfolded or if protein interactions within the kinetochore were disrupted, the connectivity between centromeric DNA and the microtubules would be compromised. The typical force required to unfold a protein or break interactions is in the range of 10–100 pN (Weisel et al., 2003; Lin et al., 2005; Kumar and Li, 2010). Nicklas did not observe an immediate rupture of chromosome–spindle attachments even while applying 700 pN on chromosomes, suggesting that the kinetochore is constructed in a way that can withstand high loads.Open in a separate windowFigure 2.Models for force response at kinetochores at both the individual protein level and global scale. (A–C) We propose three nonexclusive models for how kinetochores respond to the application of force: kinetochore proteins with elastic properties could serve to absorb some of the force produced by depolymerizing microtubules (A), multiple weak interfaces could form parallel attachments between the depolymerizing microtubule and chromosome such that the force produced by the microtubule would be diffused across multiple connections (B), and additional kinetochore components could serve as dynamic cross-linkers to diffuse force and add interactions between pairs of proteins to strengthen the protein–protein interface (C). The kinetochore protein components themselves could have multiple responses at a molecular level including that (1) under pulling forces, the bonds holding together the tertiary and secondary structure of a protein can break, causing the protein to unfold. If reversible, this would provide elastic properties, but if permanent, could lead to loss of functional kinetochore components. (2) The force generated by kinetochores is directed toward the limited number of protein–DNA interactions formed between the kinetochore proteins and the chromosome. Some tension may be relieved as the DNA wrapped around adjacent nucleosomes is pulled. This first results in the straightening out of the compact “beads on a string” structure, but with sufficient pulling force, the nucleosomes would be removed from the DNA. (3) Protein–protein interfaces held together by noncovalent bonds can break under pulling force, but the presence of additional proteins to strengthen interactions could prevent the loss of important interfaces.At kinetochores, rupture events caused by force-dependent protein unfolding or the loss of protein–protein interfaces are likely avoided at least in part through the architecture and organization of the kinetochore. Previous theoretical work on the effects of force on protein structure and protein–protein interactions has highlighted organization and arrangement as key features for facilitating force resistance (Leckband, 2000; Evans, 2001). In a “series” arrangement, bonds are organized linearly such that the full force is felt by each component. However, in a “parallel” arrangement, the force is divided over multiple attachments arranged in parallel so that the force felt by each attachment is greatly reduced. The higher order organization of the kinetochore could diffuse the microtubule-generated force over multiple attachments, significantly decreasing the force that is felt by an individual kinetochore protein molecule.Although the kinetochore clearly has evolved mechanisms to accommodate potentially large cellular forces, our understanding of the architecture and organization of a kinetochore remains limited. At the level of the minimal molecular path between a microtubule and centromeric DNA, the proteins involved appear to be connected linearly (Fig. 1; Gascoigne and Cheeseman, 2011; Gascoigne et al., 2011; Bock et al., 2012; Schleiffer et al., 2012). However, there are multiple connections formed between the centromere and a single microtubule. For example, as many as 10–20 kinetochore-localized Ndc80 complexes have been measured as associating with each microtubule in both fungi and vertebrate cells (Joglekar et al., 2006, 2008; Johnston et al., 2010; Lawrimore et al., 2011), supporting a parallel model. The complexities of these connections have proven a hurdle to devising methods to measure the force produced by microtubules on specific kinetochore components or the total force exerted on the kinetochore during normal mitotic processes.In addition to defining the forces that kinetochore proteins experience, the amount of force necessary to break a bond depends on both the loading rate (force/time) and the duration of the applied force (Merkel et al., 1999). For the kinetochore, the extended periods of force experienced during metaphase (in which sister chromatids move under force in one direction for 1–2 min; Mitchison and Salmon, 1992), as well as the rapid changes in force that occur during sister chromatid oscillations, have the potential to result in a high loading rate and extended durations of applied force. As such, it will be important to account for the way that these challenges are accommodated at kinetochores. Several calculations have estimated the power output of the grasshopper and yeast spindles (Nicklas, 1988; Bloom, 2008) and provided indirect measures for the spring constant of the kinetochore based on analysis of the chromatin spring constant during anaphase (Fisher et al., 2009). However, as a result of experimental limitations, it has not been possible to precisely determine the force constant and other key force parameters at kinetochores. Without knowledge of the force constant, it is not possible to calculate the loading rate experienced by a kinetochore. Thus, defining these parameters for kinetochores is an important area for future work.

Force at the kinetochore–DNA interface

Force also has the potential to disrupt protein–DNA interactions (Fig. 2). The kinetochore is assembled on centromeric DNA, but if the kinetochore–chromatin interface were disrupted, kinetochore function would be lost. One way in which this force could be accommodated is that the force applied through the kinetochore displaces nucleosomes in pericentric regions, alleviating the mechanical stress experienced by the kinetochore itself (Bouck and Bloom, 2007; Verdaasdonk et al., 2012). Studies of the chromatin force response in S. cerevisiae have shown that a deformation of chromatin structure occurs in the regions immediately surrounding the centromere during mitosis (Pearson et al., 2001; Bouck and Bloom, 2007) and that there is an increased turnover of nucleosomes in these surrounding regions (Verdaasdonk et al., 2012). Directed analyses have measured the force required to displace nucleosomes from DNA. These studies have obtained values of between 4 and 20 pN to irreversibly remove a nucleosome from DNA, depending on the specific approach and source of nucleosomes that was used (Cui and Bustamante, 2000; Bennink et al., 2001; Brower-Toland et al., 2002; Yan et al., 2007). For these studies, force was applied to the ends of the DNA rather than perpendicular to the DNA strand as would occur at kinetochores. This difference in the directionality of force may alter the amount of force necessary to remove a nucleosome from chromatin under mitotically applied forces.Nucleosome displacement and chromatin stretching in pericentric regions could allow the chromosome to absorb some force. However, nucleosome–DNA interactions must be maintained at the kinetochore–centromere interface. At centromeres, there are two key connections between kinetochore proteins and the underlying DNA (Gascoigne and Cheeseman, 2011; Gascoigne et al., 2011). The first occurs through the histone H3 variant, CENP-A, which epigenetically defines the centromere and forms the main site of attachment for CENP-C (Fig. 1). The other occurs via the recently identified CENP-T–W–S–X histone fold complex, which forms a heterotetrameric nucleosome-like structure (Nishino et al., 2012). Although adjacent nucleosomes surrounding the centromere could be displaced in the presence of force without severe consequences, the loss of the interaction of CENP-A or CENP-T with DNA would eliminate kinetochore function. Both the CENP-A nucleosome and the CENP-T–W–S–X complex are structurally distinct from canonical nucleosomes (Sekulic et al., 2010; Nishino et al., 2012), raising the possibility that they may have different force resistance properties. Future work characterizing the behavior of these specialized nucleosomes and the other kinetochore components will be important to understand how intrakinetochore– and kinetochore–DNA attachments are maintained in the presence of force.

How is force accommodated at kinetochores?

Although the roles of force at kinetochores have been a focus of recent work, less is known about how kinetochores are able to accommodate the forces generated at these sites. Recent work has isolated kinetochore particles from budding yeast (Akiyoshi et al., 2009) and partially reconstituted kinetochores from Xenopus laevis extract on defined templates (Guse et al., 2011). Although it is not clear how accurately these assemblies represent functional kinetochores, the reconstitution of kinetochore-like structures in vitro should allow for the analysis of its force resistance properties. At present, it remains unclear which proteins at kinetochores contribute to force resistance and how kinetochores are organized to achieve this. Current data suggest that there are two separate connections between centromeric DNA and microtubules. The first path involves an attachment of CENP-A to CENP-C followed by the Mis12 complex, which contacts KNL1 and the Ndc80 complex, with Ndc80 completing the connection to the microtubule (Fig. 1). The second connection is anchored to centromeric DNA by the CENP-T–W–S–X complex, which makes its own direct connection to the Ndc80 complex. The available biochemical data suggest that these two connections in their most minimal forms are constructed linearly and that there are two separate pools of Ndc80 that make connections to the microtubules from the Mis12 complex and CENP-T (Bock et al., 2012; Schleiffer et al., 2012; Nishino et al., 2013; Malvezzi et al., 2013). This suggests that some parts of the kinetochore might be held together by only a single protein–protein interface. However, it is possible that there are interactions between these pathways, either directly or via other protein components (Fig. 1; Gascoigne et al., 2011). If the individual protein–protein interactions within each pathway cannot withstand the force produced by the depolymerizing microtubule, the current architectural models of the kinetochore may be incomplete.Based on the currently available structural details for the kinetochore, several different models could explain how kinetochores withstand cellular forces (Fig. 2). First, kinetochore proteins may have evolved special properties that allow them to withstand force. It is possible that a subset of kinetochore proteins have elastic properties, such as those suggested by the elongation of CENP-T (Suzuki et al., 2011). Elasticity of a protein within a series arrangement would allow it to absorb some energy, thereby decreasing the force passed through the subsequent protein–protein interfaces for at least some time, much in the same way that nucleosome displacement in pericentric chromatin could diffuse the force generated at kinetochores (Verdaasdonk et al., 2012). In this model, energy is absorbed by breaking or rearranging bonds within kinetochore proteins rather than between proteins, thereby protecting the key interfaces within the kinetochore. Second, the connections between the microtubule and centromere are likely to be arranged in a parallel manner such that they sum to a strong interface. The multiple copies of each core kinetochore protein that are present per microtubule (Joglekar et al., 2006, 2008; Johnston et al., 2010; Lawrimore et al., 2011) support at least a partial contribution from this parallel model. Third, there may be additional kinetochore proteins that are not part of the linear connectivity between the centromere and microtubule but that strengthen connections between kinetochore components that would otherwise be too weak. For example, the Tetrahymena thermophila cilia protein Bld10 has recently been proposed to structurally stabilize the basal body under the force generated during cilia beating (Bayless et al., 2012). At kinetochores, proteins could serve a similar role either by serving as dynamic cross-linkers, connecting separate linear pathways, or by reinforcing existing connections by adding contacts between proteins. It is likely that the actual force resistance properties of the kinetochore complex require a combination of all three models.Work spanning the last 60 years has shown that the mitotic spindle can generate force that acts on kinetochores. The work we have summarized here provides a preliminary foundation for understanding the consequences of force at kinetochores, but the proposed models will change as more is discovered about kinetochore structure and organization. Defining the force resistance properties of the kinetochore will provide a better understanding of how it is able to function in the presence of force and the mechanisms by which it acts during chromosome segregation. As we look toward the future prospects of the field, the advances in the biophysical understanding of focal adhesions (Roca-Cusachs et al., 2012) provide an excellent blueprint for generating a detailed molecular picture of a large protein complex that functions under force. For focal adhesions, researchers have defined the pathway between the extracellular matrix and the cytoskeleton, analyzed the force response of each component along this pathway, and defined how cells use mechanosensors to signal to the cell. Achieving a similar understanding for the kinetochore will provide key insights into the function of this central cell division structure.     

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.     

Dynamics and Organization of Cortical Microtubules as Revealed by Superresolution Structured Illumination Microscopy

Plants employ acentrosomal mechanisms to organize cortical microtubule arrays essential for cell growth and differentiation. Using structured illumination microscopy (SIM) adopted for the optimal documentation of Arabidopsis (Arabidopsis thaliana) hypocotyl epidermal cells, dynamic cortical microtubules labeled with green fluorescent protein fused to the microtubule-binding domain of the mammalian microtubule-associated protein MAP4 and with green fluorescent protein-fused to the alpha tubulin6 were comparatively recorded in wild-type Arabidopsis plants and in the mitogen-activated protein kinase mutant mpk4 possessing the former microtubule marker. The mpk4 mutant exhibits extensive microtubule bundling, due to increased abundance and reduced phosphorylation of the microtubule-associated protein MAP65-1, thus providing a very useful genetic tool to record intrabundle microtubule dynamics at the subdiffraction level. SIM imaging revealed nano-sized defects in microtubule bundling, spatially resolved microtubule branching and release, and finally allowed the quantification of individual microtubules within cortical bundles. Time-lapse SIM imaging allowed the visualization of subdiffraction, short-lived excursions of the microtubule plus end, and dynamic instability behavior of both ends during free, intrabundle, or microtubule-templated microtubule growth and shrinkage. Finally, short, rigid, and nondynamic microtubule bundles in the mpk4 mutant were observed to glide along the parent microtubule in a tip-wise manner. In conclusion, this study demonstrates the potential of SIM for superresolution time-lapse imaging of plant cells, showing unprecedented details accompanying microtubule dynamic organization.Plant cell growth and differentiation depend on dynamic cortical microtubule organization mechanisms (Ehrhardt, 2008). Such mechanisms include branched microtubule formation and release (Murata et al., 2005; Nakamura et al., 2010; Fishel and Dixit, 2013), microtubule-templated microtubule growth (Chan et al., 2009), angle-of-contact microtubule bundling or catastrophe induction (Dixit and Cyr, 2004; Tulin et al., 2012), severing at microtubule crossovers (Wightman and Turner, 2007), and unique dynamic behavior between steady-state treadmilling and dynamic instability (Shaw et al., 2003).Cortical microtubule dynamics have been studied in vivo and in vitro with total internal reflection microscopy (TIRFM; Vizcay-Barrena et al., 2011), variable-angle emission microscopy (VAEM; Wan et al., 2011), spinning-disc microscopy (SD; Shaw and Lucas, 2011), and confocal laser scanning microscopy (CLSM; Shaw et al., 2003). TIRFM and VAEM provide sufficient resolution and speed but at limited depth of imaging (approximately 200 nm; Martin-Fernandez et al., 2013) and inevitably a very narrow field of view when used for in vivo studies (Mattheyses et al., 2010). Dynamic CLSM imaging suffers from field-of-view limitations while also introducing phototoxicity to the imaged sample. Furthermore, CLSM is based on a speed-to-resolution tradeoff that will necessitate computational extrapolation to bring resolution to affordable levels (Rosero et al., 2014). Finally, SD can provide sufficient depth and speed but otherwise poor resolution, owing to aberrations arising from the sample and the properties of the optics commonly used (Shaw and Ehrhardt, 2013).Microtubule research evolved concomitant with optical microscopy and the development of fluorescent proteins markers, allowing the resolution of microtubule dynamics and organization at video rates (Marc et al., 1998; Shaw and Ehrhardt, 2013). However, the bulk of plant cells organized in tissues and the optical properties of cell walls hamper microscopic observations, so that the delineation of fine details of microtubule organization still relies on laborious transmission electron microscopy (Kang, 2010).Alternatively, in vitro assays using total internal reflection (TIRFM) or Allen’s video-enhanced contrast-differential interference contrast microscopy (Allen et al., 1981) with purified components have advanced the understanding of microtubule-microtubule-associated protein (MAP) interactions while providing mechanistic insight on the function of MAP65 proteins (Tulin et al., 2012; Portran et al., 2013; Stoppin-Mellet et al., 2013), kinesin motors (Song et al., 1997), katanin-mediated microtubule severing (Stoppin-Mellet et al., 2007), and microtubule dynamics (Moore et al., 1997). However, it is explicitly acknowledged that such in vitro assays should be addressed in biologically coherent systems with physiological relevance to microtubule dynamics (Gardner et al., 2013; Zanic et al., 2013). Thus, an ideal approach would be to address microtubule dynamics in the complex cellular environment at spatiotemporal resolutions achieved by in vitro assays.Subdiffraction optical microscopy techniques allow subcellular observations below Abbe’s resolution threshold (Verdaasdonk et al., 2014), complementing the use of transmission electron microscopy. Such approaches permit dynamic subcellular tracking of appropriately tagged structures within living cells (Tiwari and Nagai, 2013). Practically, two superresolution strategies exist. The first involves patterned light illumination, allowing superresolution acquisitions by two fundamentally different methods, stimulated emission depletion (STED; Hell, 2007) and structured illumination microscopy (SIM; Gustafsson, 2000). The second interrogates the precision of fluorophore localization and includes stochastic optical reconstruction microscopy (STORM; Kamiyama and Huang, 2012) and photoactivation localization microscopy (PALM; Sengupta et al., 2012). The above regimes differ in translational and axial resolution, and their temporal efficiency depends on the size of the imaged area. SIM is probably the best compromise for superresolution live imaging, as it offers reasonable lateral resolution (approximately 100 nm; Gustafsson, 2000), which may be reduced to 50 nm (Rego et al., 2012), and sufficient depth of imaging combined with a reasonable axial resolution (approximately 200 nm). SIM allows dynamic imaging in a broader field of view than STED, at biologically meaningful rates compared with PALM and STORM (Kner et al., 2009), and with deeper imaging capacity compared with other superresolution regimes and with TIRFM/VAEM (Leung and Chou, 2011). Superresolution approaches have received limited attention in the plant cell biology field (Fitzgibbon et al., 2010; Kleine-Vehn et al., 2011), and their resolution potential during live imaging was not quantified previously.Here, high-numerical aperture (NA) objectives were combined with SIM for the acquisition and systematic quantification of subdiffraction details of cortical microtubules labeled either with GFP fused to the microtubule-binding domain of mammalian MAP4 (GFP-MBD; Marc et al., 1998) or with GFP fused to alpha tubulin6 (GFP-TUA6; Shaw et al., 2003). For such studies, wild-type plants and a mitogen-activated protein kinase4 (mpk4) mutant, exhibiting extensive microtubule bundling due to the overexpression and underphosphorylation of MAP65-1 (Beck et al., 2010), were used.     

ARHGEF17 sets the timer for retention of Mps1 at kinetochores

The kinetochore-associated kinase Mps1 controls the spindle assembly checkpoint, but the regulation of its kinetochore recruitment and activity is unclear. In this issue, Isokane et al. (2016. J. Cell Biol. http://dx.doi.org/10.1083/jcb.201408089) show that interaction with and phosphorylation of its substrate, ARHGEF17, regulates Mps1 kinetochore retention, suggesting an autoregulated, timer-like mechanism.To achieve mitotic fidelity, an elaborate mechanism called the spindle assembly checkpoint (SAC) has evolved to ensure proper chromosome–spindle attachment and alignment before anaphase. Through the action of many proteins found on centromeres and kinetochores, the SAC inhibits the anaphase promoting complex/cyclosome (APC/C) to prevent mitotic exit. The SAC must be highly tunable to rapidly respond to even a single misaligned chromosome, yet still allow mitosis to proceed in a timely manner once any alignment defects have been corrected. Mps1, a dual specificity kinase, is a core SAC protein that regulates kinetochore recruitment of other SAC proteins, including the protein kinases Bub1 and BubR1 and the APC/C inhibitor Mad2 (Maciejowski et al., 2010). Previous studies have implicated Mps1 dimerization and autoactivation (Hewitt et al., 2010; Jelluma et al., 2010), as well as the kinetochore component Ndc80/Hec1 and the Aurora B protein kinase, in targeting Mps1 to kinetochores (Saurin et al., 2011; Nijenhuis et al., 2013). The regulation of this targeting is less well understood, as are the mechanisms that ensure Mps1 is removed from kinetochores in a timely fashion. In this issue, Isokane et al. add ARHGEF17 to the list of Mps1 targeting machinery and propose that it mediates a timing mechanism that limits the duration of Mps1 activity at kinetochores.Isokane et al. (2016) became interested in ARHGEF17 after they uncovered this gene in the MitoCheck genome-wide RNAi screen for mitotic regulators (Neumann et al., 2010), but MitoCheck lacked the temporal resolution to determine the mitotic function of ARHGEF17. Using high-resolution confocal live-cell imaging, Isokane et al. (2016) showed that ARHGEF17-depleted HeLa cells exhibited accelerated mitoses with chromosome congression, biorientation, and segregation defects and performed phenotypic rescue using a mouse ARHGEF17 transgene. Consistent with a SAC function, Isokane et al. (2016) found that mitosis in ARHGEF17-depleted cells was not arrested in response to the microtubule poison nocodazole and that kinetochore localization of several core SAC proteins was deficient, including Bub1, BubR1, and Mad2 (Hewitt et al., 2010; Jelluma et al., 2010; Maciejowski et al., 2010). These functions of ARHGEF17 are conferred by its central Rho GEF domain (Rümenapp et al., 2002), which Isokane et al. (2016) found to be both necessary and sufficient for SAC activity. However, the SAC functions of ARHGEF17 are independent of its GEF activity, as the inactive mutant ARHGEF17^Y1216A (Zheng, 2001; Rümenapp et al., 2002) fully rescued the mitotic phenotypes.Because ARHGEF17 depletion phenocopied Mps1 inhibition, Isokane et al. (2016) explored a connection between the two and found that Mps1 interacts with the ARHGEF17 central domain and phosphorylates ARHGEF17 at three sites. Fluorescence cross-correlation spectroscopy (Wachsmuth et al., 2015) suggested that ARHGEF17 preferentially binds to inactive Mps1 in the cytoplasm, forming a complex required for localization of Mps1 and phosphorylation of its substrate, KNL1, at kinetochores (Yamagishi et al., 2012). ARHGEF17 localized to kinetochores independently of Mps1, but its only function in the SAC appears to be delivering Mps1 to kinetochores, as a kinetochore-tethered Mps1–CENP-B fusion bypassed the requirement for ARHGEF17 in the SAC. Interestingly, retention of the Mps1–ARHGEF17 complex at kinetochores is negatively regulated by Mps1 itself. The Mps1 inhibitor reversine increased recovery times for both Mps1 and ARHGEF17 in fluorescence recovery after photobleaching assays, indicating that Mps1 activity promotes their release from kinetochores. Based on these observations, Isokane et al. (2016) suggest an exciting model in which ARHGEF17 binding acts as a timer for retention of Mps1 at kinetochores: inactive Mps1 must form a complex with ARHGEF17 to bind to unattached/unaligned kinetochores, but once activated Mps1 limits its own retention at kinetochores by breaking apart the Mps1–ARHGEF17 complex (Fig. 1). Presumably, an individual molecule of Mps1 is retained at kinetochores for the time it takes it to phosphorylate the three sites on ARHGEF17 identified by Isokane et al. (2016).Open in a separate windowFigure 1.Proposed timer model for Mps1 retention at kinetochores. ARHGEF17 (red) binds inactive Mps1 (light blue) in the cytoplasm, and the complex binds to the outer kinetochore (gray) where Mps1 becomes activated (blue). Mps1 phosphorylates ARHGEF17 (blue circles), causing dissociation of both from kinetochores. It is still unknown where and how Mps1 is activated upon ARHGEF17 binding or where ARHGEF17 dissociates from Mps1 (blue question marks), and if and how Mps1 is inactivated or ARHGEF17 is dephosphorylated upon returning to the cytoplasm.This model will be very intriguing to the SAC field because it suggests a mechanism by which kinetochores achieve the proper amount of Mps1 at individual kinetochores to properly respond to chromosome alignment defects in a timely fashion. However, data presented in this study will have to be put into the context of other studies showing requirements for Ndc80/Hec1 and Aurora B in recruiting Mps1 to the kinetochore (Martin-Lluesma et al., 2002; Saurin et al., 2011; Nijenhuis et al., 2013; Zhu et al., 2013). Although Isokane et al. (2016) show that ARHGEF17 depletion has no effect on Ndc80 or Aurora B and demonstrate that its effect on Mps1 targeting is direct, how ARHGEF17 cooperates with Ndc80 and Aurora B remains to be determined. Perhaps, like Mps1, ARHGEF17 binds to Ndc80 and/or is regulated by Aurora B or perhaps Mps1 and ARHGEF17 cooperate by binding to different kinetochore components. The proposed timer model raises several exciting questions. The authors speculate that ARHGEF17 binding might activate Mps1, but this is not yet tested. It will also be important to determine the fates of Mps1 and ARHGEF17 once released from kinetochores. Does Mps1 remain active to support functions in the cytoplasm, or is it inactivated to participate in further cycles of kinetochore targeting and SAC activity? Is ARHGEF17 dephosphorylated to participate in further cycles and, if so, what is the phosphatase? The exciting study by Isokane et al. (2016) is just the first step toward addressing how and where Mps1 activity is regulated to in turn ensure timely activation and silencing of the SAC.     

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.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

Arabidopsis Separase Functions beyond the Removal of Sister Chromatid Cohesion during Meiosis

Separase is a capase family protease that is required for the release of sister chromatid cohesion during meiosis and mitosis. Proteolytic cleavage of the α-kleisin subunit of the cohesin complex at the metaphase-to-anaphase transition is essential for the proper segregation of chromosomes. In addition to its highly conserved role in cleaving the α-kleisin subunit, separase appears to have acquired additional diverse activities in some organisms, including involvement in mitotic and meiotic anaphase spindle assembly and elongation, interphase spindle pole body positioning, and epithelial cell reorganization. Results from the characterization of Arabidopsis (Arabidopsis thaliana) separase (ESP) demonstrated that meiotic expression of ESP RNA interference blocked the proper removal of cohesin from chromosomes and resulted in the presence of a mixture of fragmented chromosomes and intact bivalents. The presence of large numbers of intact bivalents raised the possibility that separase may also have multiple roles in Arabidopsis. In this report, we show that meiotic expression of ESP RNA interference blocks the removal of cohesin during both meiosis I and II, results in alterations in nonhomologous centromere association, disrupts the radial microtubule system after telophase II, and affects the proper establishment of nuclear cytoplasmic domains, resulting in the formation of multinucleate microspores.The proper segregation of chromosomes during mitosis and meiosis is dependent on the systematic formation and subsequent removal of sister chromatid cohesion, which is required for homologous chromosome pairing, recombination, and repair (for review, see Onn et al., 2008; Peters et al., 2008). It is also required for the pairwise alignment of chromosomes on the metaphase I spindle and for the generation of tension across centromeres, thereby ensuring their bipolar attachment. In mitosis, cohesion is maintained by the cohesin complex, which consists of four evolutionally conserved proteins: Sister Chromatid Cohesion1 (SCC1), SCC3, Structural Maintenance of Chromosome1 (SMC1), and SMC3 (for review, see Nasmyth and Haering, 2005). During meiosis, SCC1 is largely replaced by its meiotic homolog REC8.The establishment of sister chromatid cohesion in yeast involves a multistep process (Milutinovich et al., 2007) that begins during telophase of the previous cell cycle when cohesin subunits associate with the chromatin, ultimately becoming enriched at discrete loci termed cohesin-associated regions (Blat and Kleckner, 1999; Laloraya et al., 2000). Cohesion is established during S-phase in a process that requires the Chromosome Transmission Fidelity protein (Ctf7), which is also known as Eco1 (Skibbens et al., 1999; Toth et al., 1999) and involves the replication fork (Kenna and Skibbens, 2003; Lengronne et al., 2006). In budding yeast (Saccharomyces cerevisiae) and fission yeast (Schizosaccharomyces pombe), cohesin complexes remain on the chromosomes until mitotic anaphase (Uhlmann et al., 1999, 2000; Tomonaga et al., 2000). In contrast, in vertebrates, most cohesin complexes are released from the chromosomes during prophase in a separase-independent process (Waizenegger et al., 2000; Losada et al., 2002). The small fraction of cohesin that remains primarily in centromeric regions is released to start anaphase (Sumara et al., 2000). The release of chromosome cohesion at the metaphase-to-anaphase transition is triggered by the Cys protease, separase (ESP1), which specifically cleaves the α-kleisin subunit (Ciosk et al., 1998; Uhlmann et al., 1999, 2000; Buonomo et al., 2000; Hauf et al., 2001). Prior to the metaphase-to-anaphase transition, securin inhibits the protease activity of separase. At the onset of anaphase, securin is degraded by the anaphase-promoting complex/cyclosome freeing separase, which cleaves SCC1, facilitating the release of cohesion and chromosome separation (Cohen-Fix et al., 1996; Ciosk et al., 1998).Studies on the distribution of cohesin proteins during meiosis in a number of organisms, including yeast, Caenorhabditis elegans, mammals, and Arabidopsis (Arabidopsis thaliana), have shown that similar to the situation during mitosis in animal cells, a significant amount of cohesin is either removed from or redistributed on prophase chromosomes in a separase-independent process (Pasierbek et al., 2001; Cai et al., 2003; Eijpe et al., 2003; Lee et al., 2003; Yu and Koshland, 2005). The final resolution of chiasmata, formed as the result of homologous chromosome recombination, and the separation of homologous chromosomes depends on separase cleavage of the meiotic α-kleisin subunit, REC8, along chromosome arms at anaphase I (Buonomo et al., 2000; Kitajima et al., 2003). Centromeric cohesion is protected by the conserved SGO family of proteins until anaphase II when separase cleavage of REC8 facilitates the separation of sister chromatids (Rabitsch et al., 2003; Katis et al., 2004; McGuinness et al., 2005).In addition to its highly conserved role in cleaving the α-kleisin subunit, separase appears to have acquired additional diverse activities in different organisms (Queralt and Uhlmann, 2005). For example, separase plays a role in DNA repair by promoting the redistribution of cohesin complexes to sites of DNA damage during mitotic interphase in budding and fission yeast (Nagao et al., 2004; Strom et al., 2004). Separase is also important for mitotic anaphase spindle assembly and elongation (Jensen et al., 2001; Papi et al., 2005; Baskerville et al., 2008), interphase spindle pole body positioning (Nakamura et al., 2002), and spindle formation during meiosis in yeast (Buonomo et al., 2003). It is also important for the proper positioning of the centrosomes during the first asymmetric mitotic division, eggshell development in C. elegans (Siomos et al., 2001; Rappleye et al., 2002), and for epithelial cell reorganization and dynamics in Drosophila melanogaster (Pandey et al., 2005). In zebra fish, a separase mutation causes genome instability and increased susceptibility to epithelial cancer (Shepard et al., 2007).Results from the characterization of Arabidopsis separase suggested that the protein also has multiple roles in plants (Liu and Makaroff, 2006). Seeds homozygous for a T-DNA insert in Arabidopsis ESP exhibited embryo arrest at the globular stage with the endosperm exhibiting a weak titan-like phenotype. Furthermore, expression of ESP RNA interference (RNAi) from the meiosis-specific DMC1 promoter disrupted the proper removal of the SYN1 cohesin protein from chromosomes during meiosis and resulted in the presence of a mixture of fragmented chromosomes and intact bivalents. The presence of large numbers of intact bivalents led the authors to suggest that in addition to its requirement for the removal of cohesin, ESP may also be required for either the proper attachment of the kinetochores to the spindle or spindle function. These findings, along with the observations that separase appears to have multiple roles in other organisms, led us to conduct a detailed characterization of meiosis in ESP RNAi plants.In this report, we show that meiotic expression of ESP RNAi blocks the release of sister chromatid cohesion during both meiosis I and II, results in nonhomologous centromere association, disrupts the radial microtubule system (RMS) after telophase II, and affects the proper establishment of nuclear cytoplasmic domains. Unlike the large majority of plant meiotic mutants that have been characterized to date, reduction of ESP levels during meiosis leads to the formation of multinucleate microspores.     

The Cauliflower Mosaic Virus Protein P6 Forms Motile Inclusions That Traffic along Actin Microfilaments and Stabilize Microtubules

The gene VI product (P6) of Cauliflower mosaic virus (CaMV) is a multifunctional protein known to be a major component of cytoplasmic inclusion bodies formed during CaMV infection. Although these inclusions are known to contain virions and are thought to be sites of translation from the CaMV 35S polycistronic RNA intermediate, the precise role of these bodies in the CaMV infection cycle remains unclear. Here, we examine the functionality and intracellular location of a fusion between P6 and GFP (P6-GFP). We initially show that the ability of P6-GFP to transactivate translation is comparable to unmodified P6. Consequently, our work has direct application for the large body of literature in which P6 has been expressed ectopically and its functions characterized. We subsequently found that P6-GFP forms highly motile cytoplasmic inclusion bodies and revealed through fluorescence colocalization studies that these P6-GFP bodies associate with the actin/endoplasmic reticulum network as well as microtubules. We demonstrate that while P6-GFP inclusions traffic along microfilaments, those associated with microtubules appear stationary. Additionally, inhibitor studies reveal that the intracellular movement of P6-GFP inclusions is sensitive to the actin inhibitor, latrunculin B, which also inhibits the formation of local lesions by CaMV in Nicotiana edwardsonii leaves. The motility of P6 along microfilaments represents an entirely new property for this protein, and these results imply a role for P6 in intracellular and cell-to-cell movement of CaMV.Cauliflower mosaic virus (CaMV), the type member of the genus Caulimovirus, has a circular double-stranded DNA genome known to encode six open reading frames (ORFs). The gene product of ORF VI (P6) is a multifunctional protein whose ascribed functions have increased in number since its initial characterization over 20 years ago. P6 was originally described as the most abundant CaMV protein in infected plants (Odell and Howell, 1980) and was later shown to be the major constituent of amorphous, electron-dense inclusion bodies that are thought to be the sites of virion assembly (Fujisawa et al., 1967; Rubio-Huertos et al., 1968; Himmelbach et al., 1996; Cecchini et al., 1997). Indeed, despite the detection of other viral proteins in CaMV inclusions, the P6 protein on its own is capable of forming inclusion bodies (Cecchini et al., 1997; Li and Leisner, 2002; Haas et al., 2005).P6 is the major pathogenicity determinant for CaMV (Daubert et al., 1984; Baughman et al., 1988; Stratford and Covey, 1989; Zijlstra and Hohn, 1992) and was recently shown to be a suppressor of RNA silencing (Love et al., 2007). In addition, P6 also functions as an avirulence determinant, as it has been shown to be responsible for eliciting a hypersensitive response in Nicotiana edwardsonii and Datura stramonium, as well as nonnecrotic resistance in Nicotiana bigelovii and Arabidopsis (Arabidopsis thaliana) ectotype Tsu-O (Daubert et al., 1984; Schoelz et al., 1986; Wintermantel et al., 1993; Agama et al., 2002). The portion of the P6 protein recognized by plants is localized to the N-terminal third of the protein (Wintermantel et al., 1993; Palanichelvam et al., 2000; Agama et al., 2002). P6 also has a significant effect on plant metabolism, as it is responsible for down-regulating or inducing expression of several plant genes (Geri et al., 1999), including genes involved in ethylene signaling (Geri et al., 2004).Replication of CaMV involves the production of a polycistronic RNA intermediate, the 35S RNA, and P6 acts as a translational transactivator (TAV) by modifying the host translational machinery to allow for reinitiation of translation on this RNA (Ryabova et al., 2002). To carry out this function, the P6 protein physically interacts with the initiation factor eIF3 (Park et al., 2001), as well as ribosomal proteins L13 (Bureau et al., 2004), L18 (Leh et al., 2000), and L24 (Park et al., 2001). Finally, P6 is also a nucleocytoplasmic shuttle protein whose nuclear export is dependent upon a Leu-rich sequence near its N terminus, a region that is also involved in inclusion body formation (Li and Leisner, 2002; Haas et al., 2005). Although the precise role of the P6 protein''s nucleocytoplasmic shuttle function during infection remains to be elucidated, P6 does have the capacity to bind RNA (De Tapia et al., 1993; Cerritelli et al., 1998) and as such may act to control export of the 35S RNA from the nucleus to the cytoplasm, drawing the 35S RNA into the nascent P6 inclusion bodies where viral proteins are translated.Despite the recognized intracellular movement of P6 from cytoplasm to nucleus and the disparate cytoplasmic functions of this protein, factors controlling intracellular transport of P6 remain unknown. The cytoskeleton has been implicated in the intracellular trafficking of a number of plant viral proteins. For example, proteins encoded by several viruses have been found to colocalize with actin microfilaments, including the TGBp2 movement protein from Potato virus X (PVX), TGBp2 and TGBp3 from Potato mop-top virus, the Hsp70 homolog from Beet yellows virus, as well as both the movement (MP) and 126-kD proteins from Tobacco mosaic virus (TMV; McLean et al., 1995; Haupt et al., 2005; Ju et al., 2005; Liu et al., 2005; Prokhnevsky et al., 2005) In addition, inhibitor studies recently demonstrated that the intracellular trafficking of potato leafroll virus MP to the plasmodesmata (PD) is dependent upon an intact actin cytoskeleton (Vogel et al., 2007). Together, these studies suggest that the trafficking of viral proteins along actin filaments is a mechanism utilized by highly divergent RNA viruses.The only documented example of a plant viral protein found to colocalize with both microfilaments and microtubules in cells is the TMV MP (McLean et al., 1995; reviewed in Beachy and Heinlein, 2000; Lucas, 2006), which has been shown to associate with and stabilize microtubules and contains a motif thought to mimic the region of tubulin responsible for lateral junctions between microtubules (Boyko et al., 2000; Ashby et al., 2006). Interestingly, the CaMV gene II product (P2), an aphid transmission factor, was previously shown by immunoelectron microscopy to associate with microtubules in both insect and plant cells, although the significance of this interaction remains unclear (Blanc et al., 1996). In addition to these two viral proteins found to colocalize with microtubules in planta, the Hsp70 homolog from Beet yellows virus and the coat protein from PVX have both been shown to interact with microtubules in vitro (Karasev et al., 1992; Serazev et al., 2003). Evidence that the intracellular localization of grapevine fanleaf virus MP is disturbed by oryzalin, as well as the finding that the geminivirus replication protein AL1 interacts with a kinesin by yeast two-hybrid assay, may also indicate a potential association of these proteins with microtubules (Kong and Hanley-Bowdoin, 2002; Laporte et al., 2003).In this study, we utilize a fusion between the C terminus of P6 and GFP to visualize P6 inclusions in live cells. We demonstrate that the fusion of P6 with GFP does not interfere with its ability to act as a TAV. We further demonstrate that P6-GFP inclusion bodies move intracellularly and are associated with microtubules, actin microfilaments, and the endoplasmic reticulum (ER). Although P6-GFP inclusion bodies associated with microtubules appear stationary, we show that P6-GFP bodies can traffic along microfilaments and that this movement is severely reduced by treatment with the actin inhibitor latrunculin B (LatB). LatB treatment of N. edwardsonii leaves inhibits the formation of local lesions by CaMV, indicating the potential that P6 trafficking on microfilaments is necessary for CaMV cell-to-cell movement. Additionally, the association of P6-GFP inclusion bodies with microtubules prevents the disruption of microtubules by oryzalin, denoting a tight association between these two proteins. We discuss the potential role of P6 movement and cytoskeletal association in CaMV infection.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

The Fcp1-Wee1-Cdk1 axis affects spindle assembly checkpoint robustness and sensitivity to antimicrotubule cancer drugs

To grant faithful chromosome segregation, the spindle assembly checkpoint (SAC) delays mitosis exit until mitotic spindle assembly. An exceedingly prolonged mitosis, however, promotes cell death and by this means antimicrotubule cancer drugs (AMCDs), that impair spindle assembly, are believed to kill cancer cells. Despite malformed spindles, cancer cells can, however, slip through SAC, exit mitosis prematurely and resist killing. We show here that the Fcp1 phosphatase and Wee1, the cyclin B-dependent kinase (cdk) 1 inhibitory kinase, play a role for this slippage/resistance mechanism. During AMCD-induced prolonged mitosis, Fcp1-dependent Wee1 reactivation lowered cdk1 activity, weakening SAC-dependent mitotic arrest and leading to mitosis exit and survival. Conversely, genetic or chemical Wee1 inhibition strengthened the SAC, further extended mitosis, reduced antiapoptotic protein Mcl-1 to a minimum and potentiated killing in several, AMCD-treated cancer cell lines and primary human adult lymphoblastic leukemia cells. Thus, the Fcp1-Wee1-Cdk1 (FWC) axis affects SAC robustness and AMCDs sensitivity.The spindle assembly checkpoint (SAC) delays mitosis exit to coordinate anaphase onset with spindle assembly. To this end, SAC inhibits the ubiquitin ligase Anaphase-Promoting Complex/Cyclosome (APC/C) to prevent degradation of the anaphase inhibitor securin and cyclin B, the major mitotic cyclin B-dependent kinase 1 (cdk1) activator, until spindle assembly.¹ However, by yet poorly understood mechanisms, exceedingly prolonging mitosis translates into cell death induction.^{2, 3, 4, 5, 6, 7} Although mechanistic details are still missing on how activation of cell death pathways is linked to mitosis duration, prolongation of mitosis appears crucial for the ability of antimicrotubule cancer drugs (AMCDs) to kill cancer cells.^{2, 3, 4, 5, 6, 7} These drugs, targeting microtubules, impede mitotic spindle assembly and delay mitosis exit by chronically activating the SAC. Use of these drugs is limited, however, by toxicity and resistance. A major mechanism for resistance is believed to reside in the ability of cancer cells to slip through the SAC and exit mitosis prematurely despite malformed spindles, thus resisting killing by limiting mitosis duration.^{2, 3, 4, 5, 6, 7} Under the AMCD treatment, cells either die in mitosis or exit mitosis, slipping through the SAC, without or abnormally dividing.^{2, 3, 4} Cells that exit mitosis either die at later stages or survive and stop dividing or proliferate, giving rise to resistance.^{2, 3, 4} Apart from a role for p53, what dictates cell fate is still unknown; however, it appears that the longer mitosis is protracted, the higher the chances for cell death pathway activation are.^{2, 3, 4, 5, 6, 7}Although SAC is not required per se for killing,⁶ preventing SAC adaptation should improve the efficacy of AMCD by increasing mitosis duration.^{2, 3, 4, 5, 6, 7} Therefore, further understanding of the mechanisms by which cells override SAC may help to improve the current AMCD therapy. Several kinases are known to activate and sustain SAC, and cdk1 itself appears to be of primary relevance.^{1, 8, 9} By studying mitosis exit and SAC resolution, we recently reported a role for the Fcp1 phosphatase to bring about cdk1 inactivation.^{10, 11} Among Fcp1 targets, we identified cyclin degradation pathway components, such as Cdc20, an APC/C co-activator, USP44, a deubiquitinating enzyme, and Wee1.^{10, 11} Wee1 is a crucial kinase that controls the G2 phase by performing inhibitory phosphorylation of cdk1 at tyr-15 (Y15-cdk1). Wee1 is also in a feedback relationship with cdk1 itself that, in turn, can phosphorylate and inhibit Wee1 in an autoamplification loop to promote the G2-to-M phase transition.¹² At mitosis exit, Fcp1 dephosphorylated Wee1 at threonine 239, a cdk1-dependent inhibitory phosphorylation, to dampen down the cdk1 autoamplification loop, and Cdc20 and USP44, to promote APC/C-dependent cyclin B degradation.^{10, 11, 12} In this study we analysed the Fcp1 relevance in SAC adaptation and AMCD sensitivity.     

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

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

Introduction

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

Neuronal polarization and axon specification

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

Neuronal polarization, cytoskeletal dynamics, and polarized transport

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

Axon growth

Microtubule dynamics regulate axon growth.

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

Where does axon elongation take place?

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

Axon branching and presynaptic differentiation

Where do axon branches form?

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

Cytoskeleton dynamics and axon branch formation.

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

Is axon branching linked to axon elongation?

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

Regulation of axon branching by activity.

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

Stabilization and refinement of the axonal arborization.

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

GTP Is Required for the Microtubule Catastrophe-Inducing Activity of MAP200, a Tobacco Homolog of XMAP215

Widely conserved among eukaryotes, the microtubule-associated protein 215 (MAP215) family enhances microtubule dynamic instability. The family member studied most extensively, Xenopus laevis XMAP215, has been reported to enhance both assembly and disassembly parameters, although the mechanism whereby one protein can exert these apparently contradictory effects has not been clarified. Here, we analyze the activity of a plant MAP215 homolog, tobacco (Nicotiana tabacum) MAP200 on microtubule behavior in vitro. We show that, like XMAP215, MAP200 promotes both assembly and disassembly parameters, including microtubule growth rate and catastrophe frequency. When MAP200 is added to tubulin and taxol, strikingly long-coiled structures form. When GDP partially replaces GTP, the increase of catastrophe frequency by MAP200 is strongly diminished, even though this replacement stimulates catastrophe in the absence of MAP200. This implies that MAP200 induces catastrophes by a specific, GTP-requiring pathway. We hypothesize that, in the presence of MAP200, a catastrophe-prone microtubule lattice forms occasionally when elongated but nonadjacent protofilaments make lateral contacts.Microtubules switch stochastically between growth and shortening phases, a phenomenon known as dynamic instability (Mitchison and Kirschner, 1984). Switching from a growth phase to a shortening phase is an event termed catastrophe, and, conversely, switching from shortening to growth is termed rescue. Dynamic instability is essential for the function and organization of microtubule structures, allowing microtubule arrays to explore their environment and to be remodeled rapidly. Although dynamic instability can be observed in polymers created from pure tubulin, the characteristics of the phenomenon are subject to profound regulation by microtubule-associated proteins (Howard and Hyman, 2007).In the context of regulating dynamic instability, among myriad proteins, one family, microtubule-associated protein 215 (MAP215), has been studied particularly widely (Gard et al., 2004). This family has been reported to play a major role organizing microtubule structures in many species, including: Schizosaccharomyces pombe (Ohkura et al., 1988; Garcia et al., 2001), budding yeast (Saccharomyces cerevisiae; Severin et al., 2001), Caenorhabidis elegans (Matthews et al., 1998), Xenopus laevis (Tournebize et al., 2000), Homo sapiens (Gergely et al., 2003; Cassimeris and Morabito, 2004), Drosophila melanogaster (Goshima et al., 2005), Dictyostelium discoideum (Hestermann and Graf, 2004), Aspergillus nidulans (Enke et al., 2007), and Arabidopsis (Arabidopsis thaliana; Whittington et al., 2001; Kawamura and Wasteneys, 2008). In all of these organisms, the loss-of-function phenotype can be summarized as decreased microtubule length, indicating that MAP215, as a net result, promotes microtubule assembly.Further insight into the function of MAP215 has been gained from in vitro analysis. The extent of microtubule assembly and the rate of growth are substantially increased by Xenopus XMAP215 (Gard and Kirschner, 1987) as well as by the human ortholog, TOGp (Charrasse et al., 1998). However, interestingly, analyzing parameters of dynamic instability has revealed that XMAP215 not only promotes growth rate but also promotes shortening rate and catastrophe frequency (Vasquez et al., 1994). Catastrophe-inducing activity was also demonstrated by finding that XMAP215 can disassemble GMPCPP-stabilized microtubules (Shirasu-Hiza et al., 2003). Consistent with the idea that this protein can enhance shortening, GFP-XMAP215 labeled both growing and shortening microtubule ends (Brouhard et al., 2008), and the budding yeast ortholog, stu2, analyzed in vitro, promotes catastrophe frequency as its major activity (van Breugel et al., 2003). The ability of a protein to enhance microtubule growth as well as to increase the frequency of catastrophe is paradoxical, and the mechanism for MAP215''s bipolar activity remains to be demonstrated.In plants, orthologs of MAP215, identified as MICROTUBULE ORGANIZATION1 (MOR1) in Arabidopsis (Whittington et al., 2001) and as MAP200 in tobacco (Nicotiana tabacum; Yasuhara et al., 2002) are about 70% similar to their animal counterparts, indicating strong conservation. The effects of plant MAP215 on dynamic instability have not been characterized in vitro, although a net promotion of microtubule assembly has been observed for MAP200 (Hamada et al., 2004). In living cells, analysis of dynamic instability in wild-type and mor1-1 epidermal cells revealed mor1-1 mutation increases pause duration (Kawamura and Wasteneys, 2008). However with living cells it is difficult to distinguish direct effects of the protein from indirect effects caused by the plant.Here, we characterize dynamic instability in vitro as affected by MAP200. We confirm that the plant ortholog promotes growth, catastrophes, and rescues; however, we show that, when GDP partially replaces GTP, catastrophe promotion by MAP200 is suppressed more strongly than is growth. This result suggests that MAP215 induces catastrophe by a specific, GTP-dependent mechanism. We propose a model that predicts catastrophes promoted by MAP215 are mechanistically distinct from those arising from the loss of the GTP cap.