Dynamic Subunit Exchange and the Regulation of Microtubule Assembly by the Stress Response Protein Human αB Crystallin

Background

The small heat shock protein (sHSP), human αB crystallin, forms large, polydisperse complexes that modulate the tubulin-microtubule equilibrium using a dynamic mechanism that is poorly understood. The interactive sequences in αB crystallin for tubulin are surface exposed, and correspond to interactive sites for the formation of αB crystallin complexes.

Methodology/Principal Findings

There is sequence homology between tubulin and the interactive domains in the β8-strand of the core domain and the C-terminal extension of αB crystallin. This study investigated the hypothesis that the formation of tubulin and αB crystallin quaternary structures was regulated through common interactive domains that alter the dynamics of their assembly. Size exclusion chromatography (SEC), SDS-PAGE, microtubule assembly assays, aggregation assays, multiple sequence alignment, and molecular modeling characterized the dynamic response of tubulin assembly to increasing concentrations of αB crystallin. Low molar ratios of αB crystallin∶tubulin were favorable for microtubule assembly and high molar ratios of αB crystallin∶tubulin were unfavorable for microtubule assembly. Interactions between αB crystallin and unassembled tubulin were observed using SEC and SDS-PAGE.

Conclusions/Significance

Subunits of αB crystallin that exchange dynamically with the αB crystallin complex can interact with tubulin subunits to regulate the equilibrium between tubulin and microtubules.     

The contribution of αβ-tubulin curvature to microtubule dynamics

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

The cycle of microtubule polymerization.

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

Tubulin dimers and their curvatures

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

Conformation and dynamic instability

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

Microtubule depolymerases stabilize curved conformations of tubulin

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

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

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

Sensing conformation at lattice contacts

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

Conclusions and outlook

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

Cancer Cells Acquire Mitotic Drug Resistance Properties Through Beta I-Tubulin Mutations and Alterations in the Expression of Beta-Tubulin Isotypes

Background

Anti-mitotic compounds (microtubule de-stabilizers) such as vincristine and vinblastine have been shown clinically successful in treating various cancers. However, development of drug-resistance cells limits their efficacies in clinical situations. Therefore, experiments were performed to determine possible drug resistance mechanisms related to the application of anti-mitotic cancer therapy.

Principal Findings

A KB-derived microtubule de-stabilizer-resistant KB-L30 cancer cell line was generated for this study. KB-L30 cells showed cross-resistance to various microtubule de-stabilizers including BPR0L075, vincristine and colchicine through multiple-drug resistant (MDR)-independent mechanisms. Surprisingly, KB-L30 cells showed hyper-sensitivity to the microtubule-stabilizer, paclitaxel. Results of the RT-PCR analysis revealed that expression of both class II and III β-tubulin was down-regulated in KB-L30 cells as compared to its parental KB cancer cells. In addition, DNA sequencing analysis revealed six novel mutation sites present in exon four of the βI-tubulin gene. Computational modeling indicated that a direct relationship exists between βI-tubulin mutations and alteration in the microtubule assembly and dynamic instability in KB-L30 cells and this predicted model was supported by an increased microtubule assembly and reduced microtubule dynamic instability in KB-L30 cells, as shown by Western blot analysis.

Conclusions and Significance

Our study demonstrated that these novel mutations in exon four of the βI-tubulin induced resistance to microtubule de-stabilizers and hyper-sensitivity to microtubule stabilizer through an alteration in the microtubule assembly and dynamics in cancer cells. Importantly, the current study reveals that cancer cells may acquire drug resistance ability to anti-mitotic compounds through multiple changes in the microtubule networks. This study further provided molecular information in drug selection for patients with specific tubulin mutations.     

Roles of β-Tubulin Residues Ala428 and Thr429 in Microtubule Formation in Vivo

The C termini of β-tubulin isotypes are regions of high sequence variability that bind to microtubule-associated proteins and motors and undergo various post-translational modifications such as polyglutamylation and polyglycylation. Crystallographic analyses have been unsuccessful in resolving tubulin C termini. Here, we used a stepwise approach to study the role of this region in microtubule assembly. We generated a series of truncation mutants of human βI and βIII tubulin. Transient transfection of HeLa cells with the mutants shows that mutants with deletions of up to 22 residues from βIII and 16 from βI can assemble normally. Interestingly, removal of the next residue (Ala⁴²⁸) results in a complete loss of microtubule formation without affecting dimer formation. C-terminal tail switching of human βI and βIII tubulin suggests that C-terminal tails are functionally equivalent. In short, residues outside of 1–429 of human β-tubulins make no contribution to microtubule assembly. Ala⁴²⁸, in the C-terminal sequence motif N-QQYQDA⁴²⁸, lies at the end of helix H12 of β-tubulin. We hypothesize that this residue is important for maintaining helix H12 structure. Deletion of Ala⁴²⁸ may lead to unwinding of helix H12, resulting in tubulin dimers incapable of assembly. Thr⁴²⁹ plays a more complex role. In the βI isotype of tubulin, Thr⁴²⁹ is not at all necessary for assembly; however, in the βIII isotype, its presence strongly favors assembly. This result is consistent with a likely more complex function of βIII as well as with the observation that evolutionary conservation is total for Ala⁴²⁸ and frequent for Thr⁴²⁹.Microtubules are involved in a great variety of cellular functions. Their constituent protein tubulin is an αβ heterodimer, both α- and β-tubulin existing as multiple isotypes, encoded by different genes and differing in amino acid sequence (1). The differences among the isotypes are highly conserved in evolution. In mammals, the β isotypes are βIa, βIb, βII, βIII, βIVa, βIVb, βV, and βVI. There is evidence that the isotype differences have functional significance. For instance, the βIV isotype is found in all axonemes (2).Structurally, both α- and β-tubulin consist of a globular region of 427 amino acids followed by a C-terminal region of 17–24 amino acids (3–5). The C-terminal region is highly negatively charged, being especially rich in glutamate residues and lacking in basic residues, and is likely to project outward from the rest of the molecule, because of its high negative charge and the electrostatic repulsion among the glutamate residues (3). The three-dimensional structure of the globular domain has been determined by electron and x-ray crystallography (4, 5). However, the C-terminal region has never been localized in the three-dimensional reconstructions except by computer modeling. The probable reasons for this are 1) that, if the C-terminal region projects out from the rest of molecule, it is likely to be very flexible with respect to the rest of the molecule and 2) the C-terminal region undergoes post-translational modification. Both of these can lead to structural heterogeneity and cause the C terminus to be invisible to crystallographic techniques.In this work, we examine the role of the C termini of human β-tubulins to determine the minimal sequence requirement for microtubule incorporation through structure/function analyses. The human βI and βIII tubulin isotypes were utilized based on their high degree of sequence variability clustered at the C terminus (Fig. 1) and the fact that βI is broadly distributed among normal tissues, whereas βIII has a very narrow tissue distribution. These two isotypes share 92% sequence identity, with differences among these isotypes occurring in both the globular domain and the C-terminal region (1).Open in a separate windowFIGURE 1.Sequence alignment of human βIa and βIII tubulin isotypes. Human βIa and βIII tubulin isotypes were aligned with ClustalX 1.83 and processed with BioEdit. Hyphens denote identical residues between sequences.

TABLE 1

The C-terminal amino acid sequences of the human β-tubulin isotypes

A Screen for Kinetochore-Microtubule Interaction Inhibitors Identifies Novel Antitubulin Compounds

Background

Protein assemblies named kinetochores bind sister chromatids to the mitotic spindle and orchestrate sister chromatid segregation. Interference with kinetochore activity triggers a spindle checkpoint mediated arrest in mitosis, which frequently ends in cell death. We set out to identify small compounds that inhibit kinetochore-microtubule binding for use in kinetochore-spindle interaction studies and to develop them into novel anticancer drugs.

Methodology/Principal Findings

A fluorescence microscopy-based in vitro assay was developed to screen compound libraries for molecules that prevented the binding of a recombinant human Ndc80 kinetochore complex to taxol-stabilized microtubules. An active compound was identified that acted at the microtubule level. More specifically, by localizing to the colchicine-binding site in αβ-tubulin the hit compound prevented the Ndc80 complex from binding to the microtubule surface. Next, structure-activity analyses distinguished active regions in the compound and led to the identification of highly potent analogs that killed cancer cells with an efficacy equaling that of established spindle drugs.

Conclusions/Significance

The compound identified in our screen and its subsequently identified analogs represent new antitubulin chemotypes that can be synthetically developed into a novel class of antimitotic spindle drugs. In addition, they are stereochemically unique as their R- and S-isomers mimic binding of colchicine and podophyllotoxin, respectively, two antitubulin drugs that interact differently with the tubulin interface. Model-driven manipulation of our compounds promises to advance insight into how antitubulin drugs act upon tubulin. These advances in turn may lead to tailor-made colchicine site agents which would be valuable new assets to fight a variety of tumors, including those that have become resistant to the (antispindle) drugs used today.     

Identification and Characterization of a Novel Terrabacter ginsenosidimutans sp. nov. β-Glucosidase That Transforms Ginsenoside Rb1 into the Rare Gypenosides XVII and LXXV

A new β-glucosidase from a novel strain of Terrabacter ginsenosidimutans (Gsoil 3082^T) obtained from the soil of a ginseng farm was characterized, and the gene, bgpA (1,947 bp), was cloned in Escherichia coli. The enzyme catalyzed the conversion of ginsenoside Rb1 {3-O-[β-d-glucopyranosyl-(1-2)-β-d-glucopyranosyl]-20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol} to the more pharmacologically active rare ginsenosides gypenoside XVII {3-O-β-d-glucopyranosyl-20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol}, gypenoside LXXV {20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol}, and C-K [20-O-(β-d-glucopyranosyl)-20(S)-protopanaxadiol]. A BLAST search of the bgpA sequence revealed significant homology to family 3 glycoside hydrolases. Expressed in E. coli, β-glucosidase had apparent K_m values of 4.2 ± 0.8 and 0.14 ± 0.05 mM and V_max values of 100.6 ± 17.1 and 329 ± 31 μmol·min⁻¹·mg of protein⁻¹ against p-nitrophenyl-β-d-glucopyranoside and Rb1, respectively. The enzyme catalyzed the hydrolysis of the two glucose moieties attached to the C-3 position of ginsenoside Rb1, and the outer glucose attached to the C-20 position at pH 7.0 and 37°C. These cleavages occurred in a defined order, with the outer glucose of C-3 cleaved first, followed by the inner glucose of C-3, and finally the outer glucose of C-20. These results indicated that BgpA selectively and sequentially converts ginsenoside Rb1 to the rare ginsenosides gypenoside XVII, gypenoside LXXV, and then C-K. Herein is the first report of the cloning and characterization of a novel ginsenoside-transforming β-glucosidase of the glycoside hydrolase family 3.Ginseng refers to the roots of members of the plant genus Panax, which have been used as a traditional medicine in Asian countries for over 2,000 years due to their observed beneficial effects on human health. Ginseng saponins, also referred to as ginsenosides, are the major active components of ginseng (27). Various biological activities have been ascribed to ginseng saponins, including anti-inflammatory activity (43), antitumor effects (23, 39), and neuroprotective and immunoprotective (15, 31) effects.Ginsenosides can be categorized as protopanaxadiol (PPD), protopanaxatriol, and oleanane saponins, based on the structure of the aglycon, with a dammarane skeleton (29). The PPD-type ginsenosides are further classified into subgroups based on the position and number of sugar moieties attached to the aglycon at positions C-3 and C-20. For example, one of the largest PPD-type ginsenosides, Rb1 {3-O-[β-d-glucopyranosyl-(1-2)-β-d-glucopyranosyl]-20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol}, contains 4 glucose moieties, two each attached via glycosidic linkages to the C-3 and C-20 positions of the aglycon (Fig. ​(Fig.11).Open in a separate windowFIG. 1.Chemical structures of protopanaxadiol and protopanaxatriol ginsenosides (5). The ginsenosides represented here are all (S)-type ginsenosides. glc, β-d-glucopyranosyl; arap, α-l-arabinopyranosyl; araf, α-l-arabinofuranosyl; rha, α-l-rhamnopyranosyl; Gyp, gypenoside; C, compound.Because of their size, low solubility, and poor permeability across the cell membrane, it is difficult for human body to directly absorb large ginsenosides (44), although these components constitute the major portion of the total ginsenoside in raw ginseng (30). Moreover, the lack of the availability of the rare ginsensoides limits the research on their biological and medicinal properties. Therefore, transformation of these major ginsenosides into smaller deglycosylated ginsenosides, which are more effective in in vivo physiological action, is required (1, 37).The production of large amounts of rare ginsenosides from the major ginsenosides can be accomplished through a number of physiochemical methods such as heating (17), acid treatment (2), and alkali treatment (48). However, these approaches produce nonspecific racemic mixtures of rare ginsenosides. As an alternative, enzymatic methods have been explored as a way to convert the major ginsenosides into more pharmacologically active rare ginsenosides in a more specific manner (14, 20).To date, three types of glycoside hydrolases, β-d-glucosidase, α-l-arabinopyranosidase, and α-l-arabinofuranosidase, have been found to be involved in the biotransformation of PPD-type ginsenosides. For example, a β-glucosidase isolated from a fungus converts Rb1 to C-K [20-O-(β-d-glucopyranosyl)-20(S)-protopanaxadiol] (45), and an α-l-arabinopyranosidase and α-l-arabinofuranosidase have been isolated from an intestinal bacterium that hydrolyze, respectively, Rb2 {3-O-[β-d-glucopyranosyl-(1-2)-β-d-glucopyranosyl]-20-O-[α-l-arabinopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol} to Rd {3-O-[β-d-glucopyranosyl-(1-2)-β-d-glucopyranosyl]-20-O-β-d-glucopyranosyl-20(S)-protopanaxadiol} and Rc {3-O-[β-d-glucopyranosyl-(1-2)-β-d-glucopyranosyl]-20-O- [α-l-arabinofuranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol} to Rd (34). Two recombinant enzymes that convert major ginsenosides into rare ginsenosides have been cloned and expressed in Escherichia coli: Solfolobus solfataricus β-glycosidase, which transforms Rb1 or Rc to C-K (28), and β-glucosidase from a soil metagenome, which transforms Rb1 to Rd (16). Both of these glycoside hydrolases are family 1 glycoside hydrolases.Here, we report the cloning and expression in E. coli of a gene (bgpA) encoding a new ginsenoside-hydrolyzing β-glucosidase from a novel bacterial strain, Terrabacter ginsenosidimutans sp. nov. Gsoil 3082, isolated from a ginseng farm in Korea. BgpA is a family 3 glycoside hydrolase, and the recombinant enzyme employs a different enzymatic pathway from ginsenoside-hydrolyzing family 1 glycoside hydrolases. BgpA preferentially and sequentially hydrolyzed the terminal and inner glucoses at the C-3 position of ginsenoside Rb1 and then the outer glucose at the C-20 position. Thus, BgpA could be effective in the biotransformation of ginsenoside Rb1 to gypenoside (Gyp) XVII {3-O-β-d-glucopyranosyl-20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol}, Gyp LXXV {20-O-[β-d-glucopyranosyl-(1-6)-β-d-glucopyranosyl]-20(S)-protopanaxadiol}, and C-K.     

Centrosomes Isolated from Spisula solidissima Oocytes Contain Rings and an Unusual Stoichiometric Ratio of α/β Tubulin

Centrosome-dependent microtubule nucleation involves the interaction of tubulin subunits with pericentriolar material. To study the biochemical and structural basis of centrosome-dependent microtubule nucleation, centrosomes capable of organizing microtubules into astral arrays were isolated from parthenogenetically activated Spisula solidissima oocytes. Intermediate voltage electron microscopy tomography revealed that each centrosome was composed of a single centriole surrounded by pericentriolar material that was studded with ring-shaped structures ~25 nm in diameter and <25 nm in length. A number of proteins copurified with centrosomes including: (a) proteins that contained M-phase–specific phosphoepitopes (MPM-2), (b) α-, β-, and γ-tubulins, (c) actin, and (d) three low molecular weight proteins of <20 kD. γ-Tubulin was not an MPM-2 phosphoprotein and was the most abundant form of tubulin in centrosomes. Relatively little α- or β-tubulin copurified with centrosomes, and the ratio of α- to β-tubulin in centrosomes was not 1:1 as expected, but rather 1:4.6, suggesting that centrosomes contain β-tubulin that is not dimerized with α-tubulin.     

Tubulin Tyrosination Is Required for the Proper Organization and Pathfinding of the Growth Cone

Background

During development, neuronal growth cones integrate diffusible and contact guidance cues that are conveyed to both actin and microtubule (MT) cytoskeletons and ensure axon outgrowth and pathfinding. Although several post-translational modifications of tubulin have been identified and despite their strong conservation among species, their physiological roles during development, especially in the nervous sytem, are still poorly understood.

Methodology/Findings

Here, we have dissected the role of a post-translational modification of the last amino acid of the α-tubulin on axonal growth by analyzing the phenotype of precerebellar neurons in Tubulin tyrosin ligase knock-out mice (TTL ^−/−) through in vivo, ex vivo and in vitro analyses. TTL ^−/− neurons are devoid of tyrosinated tubulin. Their pathway shows defects in vivo, ex vivo, in hindbrains open-book preparations or in vitro, in a collagen matrix. Their axons still orient toward tropic cues, but they emit supernumerary branches and their growth cones are enlarged and exhibit an emission of mis-oriented filopodia. Further analysis of the TTL ^−/− growth cone intracellular organization also reveals that the respective localization of actin and MT filaments is disturbed, with a decrease in the distal accumulation of Myosin IIB, as well as a concomitant Rac1 over-activation in the hindbrain. Pharmacological inhibition of Rac1 over-activation in TTL ^−/− neurons can rescue Myosin IIB localization.

Conclusions/Significance

In the growth cone, we propose that tubulin tyrosination takes part in the relative arrangement of actin and MT cytoskeletons, in the regulation of small GTPases activity, and consequently, in the proper morphogenesis, organization and pathfinding of the growth cone during development.     

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.     

Rescue of a Dominant Mutant With RNA Interference

Maize Mucronate1 is a dominant floury mutant based on a misfolded 16-kDa γ-zein protein. To prove its function, we applied RNA interference (RNAi) as a dominant suppressor of the mutant seed phenotype. A γ-zein RNAi transgene was able to rescue the mutation and restore normal seed phenotype. RNA interference prevents gene expression. In most cases, this is used to study gene function by creating a new phenotype. Here, we use it for the opposite purpose. We use it to reverse the creation of a mutant phenotype by restoring the normal phenotype. In the case of the maize Mucronate1 (Mc1) phenotype, interaction of a misfolded protein with other proteins is believed to be the basis for the Mc1 phenotype. If no misfolded protein is present, we can reverse the mutant to the normal phenotype. One can envision using this approach to study complex traits and in gene therapy.TRANSLUCENT or vitreous maize kernels are harder and able to sustain stronger mechanical strength during harvesting, transportation, and storage. There is a direct link between a vitreous seed phenotype and the type of storage proteins in the seed, collectively called zeins in maize. Zeins, encoded by a multigene family, constitute >60% of all maize seed proteins. They are classified into four groups (α-, β-, γ-, and δ-zein) on the basis of their structures (Esen 1987). Zeins are specifically synthesized in the endosperm ∼10 days after pollination (DAP) and deposited into protein bodies (Wolf et al. 1967; Burr and Burr 1976; Lending and Larkins 1992). Irregularly shaped protein bodies are found in floury or opaque kernel phenotypes (Coleman et al. 1997; Kim et al. 2004, 2006; Wu et al. 2010; Wu and Messing 2010). The terms “floury” and “opaque” were originally created on the basis of the genetic behaviors of the mutant allele causing the soft kernel texture. The floury mutants behave as semidominant or dominant mutants, as floury1 and floury2 do, while the opaque mutants are recessive, as opaque1 and opaque2 are (Hayes and East 1915; Lindstrom 1923; Emerson et al. 1935; Maize Genetics Cooperation 1939). Similar to floury2 with a single mutation in the signal peptide of a 22-kDa α-zein resulting in an unprocessed protein (Coleman et al. 1995), De*-B30 produces an unprocessed 19-kDa α-zein (Kim et al. 2004). It was hypothesized that the two mutant proteins with an unprocessed signal peptide are misfolded and docked in the membranes of the rough endoplasmic reticulum (RER), blocking the deposition of other zein proteins (Coleman et al. 1995; Kim et al. 2004). In Mucronate1 (Mc1), a 38-bp deletion in the C terminus of the 16-kDa γ-zein (γ16-zein) gene resulted in a frameshift and a protein with a different amino-acid tail. This modified 16-kDa γ-zein (Δγ16-zein) has altered solubility properties, which would explain the formation of irregular protein bodies. Because De*-B30 and Mc1 are semidominant and dominant, respectively, they belong to the floury mutant class.The γ-zein genes (γ27-zein and γ16-zein) are homologous copies because maize underwent allotetraploidization and both gene copies have been retained during diploidization (Xu and Messing 2008). The two γ-zeins and the 15-kDa β-zein have a redundant function in stabilizing protein-body formation (Wu and Messing 2010). Knockdown of both γ-zeins with a single RNA interference (RNAi) construct conditioned only partial opacity in the crown, the top of the kernel, as opposed to the remainder or gown area of the kernel. Consistent with its light kernel phenotype, protein bodies in such a γ-zein RNAi (γRNAi) mutant exhibited a slight alteration in morphology. This phenotype is clearly distinguishable from the Mc1 phenotype, which is far more severe. Therefore, if Mc1 is caused by a misfolded chimeric 16-kDa γ-zein, preventing its expression should restore normal kernel phenotype. Indeed, a simple cross of Mc1 with a maize line carrying the γRNAi transgene produced a non-floury phenotype, providing an example of RNAi as a dominant suppressor of a dominant phenotype and as a general tool in marker rescue.

Analysis of the progeny from the cross of Mc1 and γRNAi mutants:

Mc1 seeds (Stock ID U840I) were requested from the Maize Genetics Cooperation Stock Center. The γRNAi transgenic lines have been reported in previous work (Wu et al. 2010; Wu and Messing 2010). Twelve progeny kernels from the cross of the Mc1 mutant [homozygous for the dominant-negative mutant 16-kDa γ-zein alleles (Δγ16/Δγ16) and heterozygous for the γRNAi line (γRNAi/+)] were dissected at 18 DAP for segregation and mRNA accumulation analyses. For each kernel, the embryo and endosperm were separated for DNA and RNA extraction, respectively. As shown in Figure 1A, five and seven kernels were positive and negative for the amplification of the γRNAi gene with a specific primer set, exemplifying a 1:1 segregation of the γRNAi gene.Open in a separate windowFigure 1.—Segregation analysis of the accumulations of mRNAs and proteins from the cross of the Mc1 mutant and the γRNAi line by RT–PCR and SDS–PAGE. (A) γRNAi gene segregation from progeny (Δγ16/Δγ16 x γRNAi/+) by PCR amplification with a specific primer set (GFPF, ACAACCACTACCTGAGCAC and T35SHindIII, ATTAAGCTTTGCAGGTCACTGGATTTTGG). Kernels 3, 8, 9, 10, and 12 are positive for the γRNAi gene and the rest of them are negative. M, DNA markers from top to bottom band are 3, 2, 1.5, 1.4, and 1 kb. (B) RT–PCR analysis of mRNA accumulation from the normal γ16 and mutant Δγ16 alleles in the endosperms with the genotypes corresponding to the embryos analyzed above. Total RNA was extracted by using TRIzol reagent (Invitrogen). Two micrograms of RNA was digested with DNase I (Invitrogen) and then reverse-transcribed. Twenty-five nanograms of cDNA from each of the twelve endosperms was applied for PCR (25 cycles of 30 sec, 94 °C; 30 sec, 58 °C; and 1 min, 72 °C). A specific primer set (γ16F, ATGAAGGTGCTGATCGTTGC and γ16R, TCAGTAGTAGACACCGCCG) was designed for amplification of the full-length γ16-zein coding sequence (552 bp). The lower band (514 bp) from the mutant Δγ16 allele is 38 bp shorter than that from the normal allele (552 bp). Kernels 3, 8, 9, 10, and 12 with the γRNAi gene accumulated significantly less mRNA compared to those without the γRNAi gene (kernels 1, 2, 4, 5, 6, 7, and 11). BA, hybrid of B × A lines. M, DNA markers from top to bottom are 1 kb, 750 bp, and 500 bp. (C) Profile of zein accumulations of 20 kernels from the progeny as described in the text. The zein extraction method has been described elsewhere (Wu et al. 2009). The Δγ16-zein from Mc1 was not extracted by traditional total-zein extraction protocol (70% ethanol and 2% 2-mercaptoethanol). The γ27- and γ16-zeins were knocked down to a nondetectable level in kernels 1, 2, 3, 5, 7, 10, 12, 13, 16, and 20. In γRNAi-gene segregating progeny (kernels 4, 6, 8, 9, 11, 14, 15, 17, 18, and 19), the γ16-zein from the normal γ16 allele is marked by arrowheads. Protein loaded in each lane was equal to 500 μg fresh endosperm at 18 DAP. The size for each band is indicated by the numbers in the “kDa” columns. BA, hybrid of B × A lines; 1–20, kernels from the progeny described above; M, protein markers from top to bottom are 50, 25, 20, and 15 kDa.Due to the 38-bp deletion in the C terminus of the coding region, the Δγ16 allele is shorter than the normal one (Figure 1B). Therefore, most of Δγ16-zein was in the non-zein fraction. In progeny endosperms of another 20 kernels from the same cross described above segregating for the γRNAi gene, two types of γ16-zeins were synthesized: the normal γ16-zein in the ethanol-soluble zein fraction and the Δγ16-zein in the non-zein fraction. In progeny inheriting the γRNAi gene, the γ27- and γ16-zeins were reduced to nondetectable levels (Figure 1C). Although the Δγ16-zein is not in the ethanol-soluble zein fraction, the level of normal γ16-zein is a good indicator of the accumulation of the Δγ16-zein.

Rescue of protein-body morphologies in the Mc1 mutant:

Regular protein bodies are round with distinct membrane boundaries (Figure 2A) and 1–2 μm in diameter at maturity. In homozygous and heterozygous Mc1 mutants (Δγ16/Δγ16 and Δγ16/+), protein bodies were irregularly shaped, some without discrete boundaries (Figure 2, C and D), which is quite different from the absence of normal γ27- or γ16-zeins in maize endosperm (Figure 2B). Indeed, protein bodies of the Mc1 mutant, blocked in the accumulation of Δγ16-zein, showed morphologies with no discernible difference from those in the γRNAi/+ line (Figure 2, B and E).Open in a separate windowFigure 2.—Transmission electron micrographs of protein bodies. The method has been described elsewhere (Wu and Messing 2010). (A) Nontransgenic BA. (B) γRNAi transgenic line (γRNAi/+). (C) Mc1 (Δγ16/Δγ16). (D) Cross of Mc1 mutant and nontransgenic hybrid of B × A lines (Δγ16/+). (E) Cross of Mc1 mutant (Δγ16/Δγ16) and heterologous γRNAi transgenic line (γRNAi/+). PB, protein body; RER, rough endoplasmic reticulum; CW, cell wall; Mt, mitochondria; SG, starch granule. Bars, 500 nm.

Recovery of floury phenotype in progeny:

On the basis of these observations, it is reasoned that irregularly shaped protein bodies (Figure 2, C and D) in the Mc1 mutant cause the floury phenotype (Figure 3, A and B). Because knockdown of γ-zeins caused opacity only in the crown area (Figure 3C), one could envision that once the irregular protein bodies are restored, the kernel would become vitreous in the gown area of the kernel. Indeed, the progeny ear from the cross of Δγ16/Δγ16 and γRNAi/+ showed a 1:1 ratio of floury and vitreous kernels (Figure 3, D and F), and all kernels were vitreous when the Mc1 mutant was pollinated by a homozygous γRNAi line (Figure 3E).Open in a separate windowFigure 3.—Segregation of vitreous and floury kernels from a progeny ear. (A) Mc1 mutant with Δγ16/Δγ16 genotype. (B) The cross of the Mc1 mutant and the nontransgenic hybrid of B × A lines, showing floury phenotype as in A. (C) γRNAi transgenic line with partial opacity only in the crown area. (D) The cross of the Mc1 mutant (Δγ16/Δγ16) and the heterologous γRNAi transgenic line (γRNAi/+), showing a 1:1 ratio of vitreous and floury kernels. A row in the ear is marked with arrowheads and crosses to indicate vitreous and floury gowns of kernels. (E) Cross of the Mc1 mutant (Δγ16/Δγ16) and the γRNAi homozygous transgenic line (γRNAi/γRNAi), showing all vitreous kernels. (F) Truncated kernel phenotype. (Top) Mc1, cross of Mc1 × BA, and γRNAi transgenic line. (Bottom) Three vitreous and floury kernels from D.

Conclusions:

RNAi can be used to rescue mutations that are dominant negative with a single cross, providing a useful tool in genetic analysis, plant breeding, and potentially in gene therapy in general.     

Genetic evidence for interaction between eta- and beta-tubulins

The thermosensitive allelic mutations sm19-1 and sm19-2 of Paramecium tetraurelia cause defective basal body duplication: growth at the nonpermissive temperature yields smaller and smaller cells with fewer and fewer basal bodies. Complementation cloning of the SM19 gene identified a new tubulin, eta-tubulin, showing low homology with each of the other five tubulins, α to , characterized in P. tetraurelia. In order to analyze η-tubulin functions, we used a genetic approach to identify interacting molecules. Among a series of extragenic suppressors of the sm19-1 mutation, the su3-1 mutation was characterized as an E288K substitution in the β-PT2 gene coding for a β-tubulin, while the mutation noc^r1 conferring nocodazole resistance and localized in another β-tubulin gene, β-PT3, was shown to enhance the mutant phenotype. The interaction between η-tubulin and microtubules, revealed by genetic data, is supported by two further types of evidence: first, the mutant phenotype is rescued by taxol, which stabilizes microtubules; second, molecular modeling suggests that η-tubulin, like γ- and δ-tubulins, might be a microtubule minus-end capping molecule. The likely function of η-tubulin as part of a complex specifically involved in basal body biogenesis is discussed.     

Acetylation/deacetylation and microtubule associated proteins influence flagellar axonemal stability and sperm motility

PTMs and microtubule-associated proteins (MAPs) are known to regulate microtubule dynamicity in somatic cells. Reported literature on modulation of α-tubulin acetyl transferase (αTAT1) and histone deacetylase 6 (HDAC6) in animal models and cell lines illustrate disparity in correlating tubulin acetylation status with stability of MT. Our earlier studies showed reduced acetyl tubulin in sperm of asthenozoospermic individuals. Our studies on rat sperm showed that on inhibition of HDAC6 activity, although tubulin acetylation increased, sperm motility was reduced. Studies were therefore undertaken to investigate the influence of tubulin acetylation/deacetylation on MT dynamicity in sperm flagella using rat and human sperm. Our data on rat sperm revealed that HDAC6 specific inhibitor Tubastatin A (T) inhibited sperm motility and neutralized the depolymerizing and motility debilitating effect of Nocodazole. The effect on polymerization was further confirmed in vitro using pure MT and recHDAC6. Also polymerized axoneme was less in sperm of asthenozoosperm compared to normozoosperm. Deacetylase activity was reduced in sperm lysates and axonemes exposed to T and N+T but not in axonemes of sperm treated similarly suggesting that HDAC6 is associated with sperm axonemes or MT. Deacetylase activity was less in asthenozoosperm. Intriguingly, the expression of MDP3 physiologically known to bind to HDAC6 and inhibit its deacetylase activity remained unchanged. However, expression of acetyl α-tubulin, HDAC6 and microtubule stabilizing protein SAXO1 was less in asthenozoosperm. These observations suggest that MAPs and threshold levels of MT acetylation/deacetylation are important for MT dynamicity in sperm and may play a role in regulating sperm motility.     

C-Terminal Tail Polyglycylation and Polyglutamylation Alter Microtubule Mechanical Properties

Microtubules are biopolymers that perform diverse cellular functions. Microtubule behavior regulation occurs in part through post-translational modification of both the α- and β-subunits of tubulin. One class of modifications is the heterogeneous addition of glycine and/or glutamate residues to the disordered C-terminal tails (CTTs) of tubulin. Because of their prevalence in stable, high-stress cellular structures such as cilia, we sought to determine if these modifications alter microtubules’ intrinsic stiffness. Here, we describe the purification and characterization of differentially modified pools of tubulin from Tetrahymena thermophila. We found that post-translational modifications do affect microtubule stiffness but do not affect the number of protofilaments incorporated into microtubules. We measured the spin dynamics of nuclei in the CTT backbone by NMR spectroscopy to explore the mechanism of this change. Our results show that the α-tubulin CTT does not protrude out from the microtubule surface, as is commonly depicted in models, but instead interacts with the dimer’s surface. This suggests that the interactions of the α-tubulin CTT with the tubulin body contributes to the stiffness of the assembled microtubule, thus providing insight into the mechanism by which polyglycylation and polyglutamylation can alter microtubule mechanical properties.     

Asparagine synthetase regulates lung-cancer metastasis by stabilizing the β-catenin complex and modulating mitochondrial response

The availability of asparagine is the limitation of cell growth and metastasis. Asparagine synthetase (ASNS) was an essential enzyme for endogenous asparagine products. In our study, ASNS-induced asparagine products were essential to maintain tumor growth and colony formations in vitro. But mutated ASNS which defected endogenous asparagine products still upregulated cell invasiveness, which indicated that ASNS promoted invasiveness by alternative pathways. Mechanically, ASNS modulated Wnt signal transduction by promoting GSK3β phosphorylation on ser9 and stabilizing the β-catenin complex, as result, ASNS could promote more β-catenin translocation into nucleus independent of endogenous asparagine. At the same time, ASNS modulated mitochondrial response to Wnt stimuli with increased mitochondrial potential and membrane fusion. In summary, ASNS promoted metastasis depending on Wnt pathway and mitochondrial functions even without endogenous asparagine products.Subject terms: Non-small-cell lung cancer, Cell invasion, Hydrolases

In the Wnt pathway, phosphorylation of GSK3β(S9) is very important to stabilize β-catenin complex, ASNS can upregulate phosphorylation of AKT on Ser473, then promote GSK3β phosphorylation on Ser9 to stabilize β-catenin complex. When Wnt pathway is activated, ASNS elevates p-DRP1(637) and decreases p-DRP1(616) to suppress mitochondrial fission.      

The Effects of Orally Administered Beta-Glucan on Innate Immune Responses in Humans,a Randomized Open-Label Intervention Pilot-Study

Rationale

To prevent or combat infection, increasing the effectiveness of the immune response is highly desirable, especially in case of compromised immune system function. However, immunostimulatory therapies are scarce, expensive, and often have unwanted side-effects. β-glucans have been shown to exert immunostimulatory effects in vitro and in vivo in experimental animal models. Oral β-glucan is inexpensive and well-tolerated, and therefore may represent a promising immunostimulatory compound for human use.

Methods

We performed a randomized open-label intervention pilot-study in 15 healthy male volunteers. Subjects were randomized to either the β -glucan (n = 10) or the control group (n = 5). Subjects in the β-glucan group ingested β-glucan 1000 mg once daily for 7 days. Blood was sampled at various time-points to determine β-glucan serum levels, perform ex vivo stimulation of leukocytes, and analyze microbicidal activity.

Results

β-glucan was barely detectable in serum of volunteers at all time-points. Furthermore, neither cytokine production nor microbicidal activity of leukocytes were affected by orally administered β-glucan.

Conclusion

The present study does not support the use of oral β-glucan to enhance innate immune responses in humans.

Trial Registration

ClinicalTrials.gov {"type":"clinical-trial","attrs":{"text":"NCT01727895","term_id":"NCT01727895"}}NCT01727895     

Cloning and High-Level Expression of α-Galactosidase cDNA from Penicillium purpurogenum

The cDNA coding for Penicillium purpurogenum α-galactosidase (αGal) was cloned and sequenced. The deduced amino acid sequence of the α-Gal cDNA showed that the mature enzyme consisted of 419 amino acid residues with a molecular mass of 46,334 Da. The derived amino acid sequence of the enzyme showed similarity to eukaryotic αGals from plants, animals, yeasts, and filamentous fungi. The highest similarity observed (57% identity) was to Trichoderma reesei AGLI. The cDNA was expressed in Saccharomyces cerevisiae under the control of the yeast GAL10 promoter. Almost all of the enzyme produced was secreted into the culture medium, and the expression level reached was approximately 0.2 g/liter. The recombinant enzyme purified to homogeneity was highly glycosylated, showed slightly higher specific activity, and exhibited properties almost identical to those of the native enzyme from P. purpurogenum in terms of the N-terminal amino acid sequence, thermoactivity, pH profile, and mode of action on galacto-oligosaccharides.α-Galactosidase (αGal) (EC 3.2.1.22) is of particular interest in view of its biotechnological applications. αGal from coffee beans demonstrates a relatively broad substrate specificity, cleaving a variety of terminal α-galactosyl residues, including blood group B antigens on the erythrocyte surface. Treatment of type B erythrocytes with coffee bean αGal results in specific removal of the terminal α-galactosyl residues, thus generating serological type O erythrocytes (8). Cyamopsis tetragonoloba (guar) αGal effectively liberates the α-galactosyl residue of galactomannan. Removal of a quantitative proportion of galactose moieties from guar gum by αGal improves the gelling properties of the polysaccharide and makes them comparable to those of locust bean gum (18). In the sugar beet industry, αGal has been used to increase the sucrose yield by eliminating raffinose, which prevents normal crystallization of beet sugar (28). Raffinose and stachyose in beans are known to cause flatulence. αGal has the potential to alleviate these symptoms, for instance, in the treatment of soybean milk (16).αGals are also known to occur widely in microorganisms, plants, and animals, and some of them have been purified and characterized (5). Dey et al. showed that αGals are classified into two groups based on their substrate specificity. One group is specific for low-M_r α-galactosides such as pNPGal (p-nitrophenyl-α-d-galactopyranoside), melibiose, and the raffinose family of oligosaccharides. The other group of αGals acts on galactomannans and also hydrolyzes low-M_r substrates to various extents (6).We have studied the substrate specificity of αGals by using galactomanno-oligosaccharides such as Gal³Man₃ (6³-mono-α-d-galactopyranosyl-β-1,4-mannotriose) and Gal³Man₄ (6³-mono-α-d-galactopyranosyl-β-1,4-mannotetraose). The structures of these galactomanno-oligosaccharides are shown in Fig. ​Fig.1.1. Mortierella vinacea αGal I (11) and yeast αGals (29) are specific for the Gal³Man₃ having an α-galactosyl residue (designated the terminal α-galactosyl residue) attached to the O-6 position of the nonreducing end mannose of β-1,4-mannotriose. On the other hand, Aspergillus niger 5-16 αGal (12) and Penicillium purpurogenum αGal (25) show a preference for the Gal³Man₄ having an α-galactosyl residue (designated the stubbed α-galactosyl residue) attached to the O-6 position of the third mannose from the reducing end of β-1,4-mannotetraose. The M. vinacea αGal II (26) acts on both substrates to almost equal extents. The difference in specificity may be ascribed to the tertiary structures of these enzymes. Open in a separate windowFIG. 1Structures of galactomanno-oligosaccharides.Genes encoding αGals have been cloned from various sources, including humans (3), plants (20, 32), yeasts (27), filamentous fungi (4, 17, 24, 26), and bacteria (1, 2, 15). αGals from eukaryotes show a considerable degree of similarity and are grouped into family 27 (10).Here we describe the cloning of P. purpurogenum αGal cDNA, its expression in Saccharomyces cerevisiae, and the purification and characterization of the recombinant enzyme.     

Synthesis and bioactivities evaluation of oleanolic acid oxime ester derivatives as α-glucosidase and α-amylase inhibitors

Different oleanolic acid (OA) oxime ester derivatives (3a-3t) were designed and synthesised to develop inhibitors against α-glucosidase and α-amylase. All the synthesised OA derivatives were evaluated against α-glucosidase and α-amylase in vitro. Among them, compound 3a showed the highest α-glucosidase inhibition with an IC₅₀ of 0.35 µM, which was ∼1900 times stronger than that of acarbose, meanwhile compound 3f exhibited the highest α-amylase inhibitory with an IC₅₀ of 3.80 µM that was ∼26 times higher than that of acarbose. The inhibition kinetic studies showed that the inhibitory mechanism of compounds 3a and 3f were reversible and mixed types towards α-glucosidase and α-amylase, respectively. Molecular docking studies analysed the interaction between compound and two enzymes, respectively. Furthermore, cytotoxicity evaluation assay demonstrated a high level of safety profile of compounds 3a and 3f against 3T3-L1 and HepG2 cells.

Highlights

1. Oleanolic acid oxime ester derivatives (3a–3t) were synthesised and screened against α-glucosidase and α-amylase.
2. Compound 3a showed the highest α-glucosidase inhibitory with IC50 of 0.35 µM.
3. Compound 3f presented the highest α-amylase inhibitory with IC50 of 3.80 µM.
4. Kinetic studies and in silico studies analysed the binding between compounds and α-glucosidase or α-amylase.