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.     

The nuclease activity of DNA2 promotes exonuclease 1–independent mismatch repair

The DNA mismatch repair (MMR) system is a major DNA repair system that corrects DNA replication errors. In eukaryotes, the MMR system functions via mechanisms both dependent on and independent of exonuclease 1 (EXO1), an enzyme that has multiple roles in DNA metabolism. Although the mechanism of EXO1-dependent MMR is well understood, less is known about EXO1-independent MMR. Here, we provide genetic and biochemical evidence that the DNA2 nuclease/helicase has a role in EXO1-independent MMR. Biochemical reactions reconstituted with purified human proteins demonstrated that the nuclease activity of DNA2 promotes an EXO1-independent MMR reaction via a mismatch excision-independent mechanism that involves DNA polymerase δ. We show that DNA polymerase ε is not able to replace DNA polymerase δ in the DNA2-promoted MMR reaction. Unlike its nuclease activity, the helicase activity of DNA2 is dispensable for the ability of the protein to enhance the MMR reaction. Further examination established that DNA2 acts in the EXO1-independent MMR reaction by increasing the strand-displacement activity of DNA polymerase δ. These data reveal a mechanism for EXO1-independent mismatch repair.

The mismatch repair (MMR) system has been conserved from bacteria to humans (1, 2). It promotes genome stability by suppressing spontaneous and DNA damage-induced mutations (1, 3, 4, 5, 6, 7, 8, 9, 10, 11). The key function of the MMR system is the correction of DNA replication errors that escape the proofreading activities of replicative DNA polymerases (1, 4, 5, 6, 7, 8, 9, 10, 12). In addition, the MMR system removes mismatches formed during strand exchange in homologous recombination, suppresses homeologous recombination, initiates apoptosis in response to irreparable DNA damage caused by several anticancer drugs, and contributes to instability of triplet repeats and alternative DNA structures (1, 4, 5, 7, 8, 9, 10, 11, 13, 14, 15, 16, 17, 18). The principal components of the eukaryotic MMR system are MutSα (MSH2-MSH6 heterodimer), MutLα (MLH1-PMS2 heterodimer in humans and Mlh1-Pms1 heterodimer in yeast), MutSβ (MSH2-MSH3 heterodimer), proliferating cell nuclear antigen (PCNA), replication factor C (RFC), exonuclease 1 (EXO1), RPA, and DNA polymerase δ (Pol δ). Loss-of-function mutations in the MSH2, MLH1, MSH6, and PMS2 genes of the human MMR system cause Lynch and Turcot syndromes, and hypermethylation of the MLH1 promoter is responsible for ∼15% of sporadic cancers in several organs (19, 20). MMR deficiency leads to cancer initiation and progression via a multistage process that involves the inactivation of tumor suppressor genes and action of oncogenes (21).MMR occurs behind the replication fork (22, 23) and is a major determinant of the replication fidelity (24). The correction of DNA replication errors by the MMR system increases the replication fidelity by ∼100 fold (25). Strand breaks in leading and lagging strands as well as ribonucleotides in leading strands serve as signals that direct the eukaryotic MMR system to remove DNA replication errors (26, 27, 28, 29, 30). MMR is more efficient on the lagging than the leading strand (31). The substrates for MMR are all six base–base mismatches and 1 to 13-nt insertion/deletion loops (25, 32, 33, 34). Eukaryotic MMR commences with recognition of the mismatch by MutSα or MutSβ (32, 34, 35, 36). MutSα is the primary mismatch-recognition factor that recognizes both base–base mismatches and small insertion/deletion loops whereas MutSβ recognizes small insertion/deletion loops (32, 34, 35, 36, 37). After recognizing the mismatch, MutSα or MutSβ cooperates with RFC-loaded PCNA to activate MutLα endonuclease (38, 39, 40, 41, 42, 43). The activated MutLα endonuclease incises the discontinuous daughter strand 5′ and 3′ to the mismatch. A 5'' strand break formed by MutLα endonuclease is utilized by EXO1 to enter the DNA and excise a discontinuous strand portion encompassing the mismatch in a 5''→3′ excision reaction stimulated by MutSα/MutSβ (38, 44, 45). The generated gap is filled in by the Pol δ holoenzyme, and the nick is ligated by a DNA ligase (44, 46, 47). DNA polymerase ε (Pol ε) can substitute for Pol δ in the EXO1-dependent MMR reaction, but its activity in this reaction is much lower than that of Pol δ (48). Although MutLα endonuclease is essential for MMR in vivo, 5′ nick-dependent MMR reactions reconstituted in the presence of EXO1 are MutLα-independent (44, 47, 49).EXO1 deficiency in humans does not seem to cause significant cancer predisposition (19). Nevertheless, it is known that Exo1^-/- mice are susceptible to the development of lymphomas (50). Genetic studies in yeast and mice demonstrated that EXO1 inactivation causes only a modest defect in MMR (50, 51, 52, 53). In agreement with these genetic studies, a defined human EXO1-independent MMR reaction that depends on the strand-displacement DNA synthesis activity of Pol δ holoenzyme to remove the mismatch was reconstituted (54). Furthermore, an EXO1-independent MMR reaction that occurred in a mammalian cell extract system without the formation of a gapped excision intermediate was observed (54). Together, these findings implicated the strand-displacement activity of Pol δ holoenzyme in EXO1-independent MMR.In this study, we investigated DNA2 in the context of MMR. DNA2 is an essential multifunctional protein that has nuclease, ATPase, and 5''→3′ helicase activities (55, 56, 57). Previous research ascertained that DNA2 removes long flaps during Okazaki fragment maturation (58, 59, 60), participates in the resection step of double-strand break repair (61, 62, 63), initiates the replication checkpoint (64), and suppresses the expansions of GAA repeats (65). We have found in vivo and in vitro evidence that DNA2 promotes EXO1-independent MMR. Our data have indicated that the nuclease activity of DNA2 enhances the strand-displacement activity of Pol δ holoenzyme in an EXO1-independent MMR reaction.     

CXCR4 and CXCR7 transduce through mTOR in human renal cancer cells

Treatment of metastatic renal cell carcinoma (mRCC) has improved significantly with the advent of agents targeting the mTOR pathway, such as temsirolimus and everolimus. However, their efficacy is thought to be limited by feedback loops and crosstalk with other pathways leading to the development of drug resistance. As CXCR4–CXCL12–CXCR7 axis has been described to have a crucial role in renal cancer; the crosstalk between the mTOR pathway and the CXCR4–CXCL12–CXCR7 chemokine receptor axis has been investigated in human renal cancer cells. In SN12C and A498, the common CXCR4–CXCR7 ligand, CXCL12, and the exclusive CXCR7 ligand, CXCL11, activated mTOR through P70S6K and 4EBP1 targets. The mTOR activation was specifically inhibited by CXCR4 antagonists (AMD3100, anti-CXCR4-12G5 and Peptide R, a newly developed CXCR4 antagonist) and CXCR7 antagonists (anti-CXCR7-12G8 and CCX771, CXCR7 inhibitor). To investigate the functional role of CXCR4, CXCR7 and mTOR in human renal cancer cells, both migration and wound healing were evaluated. SN12C and A498 cells migrated toward CXCL12 and CXCL11; CXCR4 and CXCR7 inhibitors impaired migration and treatment with mTOR inhibitor, RAD001, further inhibited it. Moreover, CXCL12 and CXCL11 induced wound healing while was impaired by AMD3100, the anti CXCR7 and RAD001. In SN12C and A498 cells, CXCL12 and CXCL11 promoted actin reorganization characterized by thin spikes at the cell periphery, whereas AMD3100 and anti-CXCR7 impaired CXCL12/CXCL11-induced actin polymerization, and RAD001 treatment further reduced it. In addition, when cell growth was evaluated in the presence of CXCL12, CXCL11 and mTOR inhibitors, an additive effect was demonstrated with the CXCR4, CXCR7 antagonists and RAD001. RAD001-resistant SN12C and A498 cells recovered RAD001 sensitivity in the presence of CXCR4 and CXCR7 antagonists. In conclusion, the entire axis CXCR4–CXCL12–CXCR7 regulates mTOR signaling in renal cancer cells offering new therapeutic opportunities and targets to overcome resistance to mTOR inhibitors.Renal cell carcinoma (RCC) is the most lethal malignancy among urological cancers with a total of 64 770 new cases and 13 570 deaths estimated in the United States in 2012.¹ A growing understanding of the molecular biology of RCC changed the therapeutic approach toward target-based agents. Since 2005, the US Food and Drug Administration (FDA) has approved six new target agents for metastatic RCC that antagonize two principal signaling pathways: the vascular endothelial growth factor receptor (VEGF) and the mammalian target of rapamycin (mTOR).² The mTOR is an atypical intracellular serine/threonine protein kinase regulated by phosphatidylinositol 3-kinase (PI3K).³ mTOR exists in two distinct complexes termed mTOR complex 1 (mTORC1) comprising mTOR, mLST8 (also termed G-protein β-subunit-like protein, GβL, a yeast homolog of LST8), raptor (regulatory associated protein of mTOR) and PRAS40 (proline-rich Akt substrate, 40 kDa), and mTOR complex 2 (mTORC2) comprising mTOR, mLST8, rictor (rapamycin-insensitive companion of mTOR), mSin1 (mammalian stress-activated protein kinase (SAPK)-interacting protein 1), protor (protein observed with rictor) and PRR5 (proline-rich protein 5).⁴ mTORC1 responds to amino acids, stress, oxygen, energy and growth factors and is sensitive to rapamycin; when active, mTORC1 promotes cell growth and also drives cell-cycle progression. Alternatively, mTORC2 regulates cytoskeletal organization and cell survival/metabolism and is sensitive to rapamycin over longer incubation times or at higher doses.³ mTORC1 controls cell growth and translation through the phosphorylation of ribosomal protein S6 kinase (S6K) and of eukaryotic translation initiation factor 4EBP1, which regulate either the translation of ribosomal proteins or the cap-dependent translation by inhibition of eukaryotic translation initiation factor 4E, respectively.^{3, 4} The activated mTOR pathway has been identified in several human malignancies, thus being an attractive target for anticancer therapy. mTORC1 activity is inhibited by rapalogs such as rapamycin (sirolimus) and associated analogs (temsirolimus/CCI-779, RAD001, ridaforolimus/AP23573).⁵ These drugs suppress mTORC1 activity forming a complex with FK506-binding protein 12. Temsirolimus (rapamycin analog) was the first mTOR inhibitor approved as first-line treatment in patients with poor-prognosis metastatic RCC (mRCC) patients,³ ridaforolimus is currently tested in phase III clinical trials⁵ and RAD001 is indicated as second-line treatment in patients with RCC at failure of first-line treatment with sunitinib or sorafenib. Other indications are subependymal giant cell astrocytoma associated with tuberous sclerosis and progressive neuroendocrine tumors of pancreatic origin.⁵ Although mTOR inhibitors prolong progression-free survival in patients with advanced RCC, most patients develop resistance to mTOR-inhibiting agents, limiting their efficacy; the new frontier of inhibiting the mTOR pathway is to identify agents targeting the feedback loops and crosstalks with other pathways involved in the acquired resistance to mTOR inhibitors.⁶Chemokines and their receptors have been implicated in regulating RCC growth, angiogenesis and metastases.⁷ In RCC, VHL mutation resulted in HIF-dependent CXCR4 activation⁸ and CXCR4 expression predicted poor tumor-specific survival.^{8, 9, 10} Recently, CXCL12 was shown to bind with high affinity the orphan receptor CXCR7/RDC1, which also binds a second ligand in the form of interferon-inducible T-cell α chemoattractant (I-TAC/CXCL11).¹¹ Whereas the CXCR4 activity is primarily G-protein-mediated, CXCR7 is considered an atypical GPCR because ligand binding does not result in intracellular Ca2+ release.¹¹ Some studies provided evidence that CXCR7 represents a ‘decoy'' receptor, which is responsible for either sequestering extracellular CXCL12¹² or modulating CXCR4 signaling by forming CXCR7–CXCR4 heterodimers.¹³ In contrast, others demonstrated that CXCR7 relays intracellular signals^{14, 15, 16, 17} and promotes cell motility^{18, 13, 19} acting through β-arrestin.^{20, 21} CXCR7 is highly expressed in human cancers such as prostate, lung, glioma, ovarian, breast cancer cells and in tumor-associated blood vessels and seems to be essential for survival, adhesion and growth of tumor cells.^{11, 14, 15, 22, 23, 24} It was recently demonstrated that CXCR4 and CXCR7 predict prognosis in RCC.^{10, 25} CXCL12 activates CXCR4 and the derived signaling can transduce on the mTOR pathway in pancreatic cancer, gastric cancer and T-cell leukemia cells;^{26, 27, 28, 29} antagonists targeting PI3K and/or mTOR inhibited CXCL12-mediated cell migration and this effect was primarily attributed to the inhibition of mTORC1 and consequent decrease in RhoA, Cdc42 and Rac1 in human gastric carcinoma cells.²⁸Aim of the study was to evaluate interactions between the CXCL12–CXCR4–CXCR7 axis and the mTOR pathway in human renal cancer cells to identify new therapeutic opportunities and overcome resistance mechanisms.     

Flexible open conformation of the AP-3 complex explains its role in cargo recruitment at the Golgi

Vesicle formation at endomembranes requires the selective concentration of cargo by coat proteins. Conserved adapter protein complexes at the Golgi (AP-3), the endosome (AP-1), or the plasma membrane (AP-2) with their conserved core domain and flexible ear domains mediate this function. These complexes also rely on the small GTPase Arf1 and/or specific phosphoinositides for membrane binding. The structural details that influence these processes, however, are still poorly understood. Here we present cryo-EM structures of the full-length stable 300 kDa yeast AP-3 complex. The structures reveal that AP-3 adopts an open conformation in solution, comparable to the membrane-bound conformations of AP-1 or AP-2. This open conformation appears to be far more flexible than AP-1 or AP-2, resulting in compact, intermediate, and stretched subconformations. Mass spectrometrical analysis of the cross-linked AP-3 complex further indicates that the ear domains are flexibly attached to the surface of the complex. Using biochemical reconstitution assays, we also show that efficient AP-3 recruitment to the membrane depends primarily on cargo binding. Once bound to cargo, AP-3 clustered and immobilized cargo molecules, as revealed by single-molecule imaging on polymer-supported membranes. We conclude that its flexible open state may enable AP-3 to bind and collect cargo at the Golgi and could thus allow coordinated vesicle formation at the trans-Golgi upon Arf1 activation.

Eukaryotic cells have membrane-enclosed organelles, which carry out specialized functions, including compartmentalized biochemical reactions, metabolic channeling, and regulated signaling, inside a single cell. The transport of proteins, lipids, and other molecules between these organelles is mediated largely by small vesicular carriers that bud off at a donor compartment and fuse with the target membrane to deliver their cargo. The generation of these vesicles has been subject to extensive studies and has led to the identification of numerous coat proteins that are required for their formation at different sites (1, 2). Coat proteins can be monomers, but in most cases, they consist of several proteins, which form a heteromeric complex.Heterotetrameric adapter protein (AP) complexes are required at several endomembranes for cargo binding. Five well-conserved AP-complexes with differing functions have been identified in mammalian cells, named AP-1–AP-5, of which three (AP-1–AP-3) are conserved from yeast to human (3, 4). The three conserved adapter complexes function at different membranes along the endomembrane system. AP-1 is required for cargo transport between the Golgi and the endosome, AP-2 is required for cargo recognition and transport between the plasma membrane and the early endosome. Finally, AP-3 functions between the trans Golgi and the vacuole in yeast, whereas mammalian AP-3 localizes to a tubular endosomal compartment, in addition to or instead of the TGN (2, 5, 6).Each of the complexes consists of four different subunits: two large adaptins (named α−ζ and β1-5 respectively), a medium-sized subunit (μ1-5), and a small subunit (σ1-5). While μ- and σ-subunits together with the N-termini of the large adaptins build the membrane-binding core of the complex, the C-termini of both adaptins contain the ear domains, which are connected via flexible linkers (2). The recruitment of these complexes to membranes is not entirely conserved. They all require cargo binding, yet AP-1 binds Arf1-GTP with the γ and β1 subunit and phosphatidylinositol-4-phosphate (PI4P) via a proposed conserved site on its γ-subunit (7, 8). AP-2, on the other hand, interacts with PI(4,5)P₂ at the plasma membrane via its α, β2, and μ2 subunits (9, 10, 11).Several studies have uncovered how AP-3 functions in cargo sorting in yeast. AP-3 recognizes cargo at the Golgi via two sorting motifs in the cytosolic segments of membrane proteins: a Yxxφ sorting motif, as found in yeast in the SNARE Nyv1 or the Yck3 casein kinase, which binds to a site in μ3, as shown for mammalian AP-3, which is similar to μ2 in AP-2 (12, 13, 14), and dileucine motifs as found in the yeast SNARE Vam3 or the alkaline phosphatase Pho8, potentially also at a site comparable to AP-1 and AP-2 (15, 16). Unlike AP-1 and AP-2-coated vesicles, which depend on clathrin for their formation (2, 17), AP-3 vesicle formation in yeast does not require clathrin or the HOPS subunit Vps41 (18), yet Vps41 is required at the vacuole to bind AP-3 vesicles prior to fusion (19, 20, 21, 22). Studies in metazoan cells revealed that Vps41 and AP-3 function in regulated secretion (23, 24, 25), and AP-3 is required for biogenesis of lysosome-related organelles (26). This suggests that the AP-3 complex has features that are quite different from AP-1 and AP-2 complexes, which cooperate with clathrin in vesicle formation (2).Among the three conserved AP complexes, the function of the AP-3 complex is the least understood. Arf1 is necessary for efficient AP-3 vesicle generation in mammalian cells and shows a direct interaction with the β3 and δ subunits of AP-3 (27, 28). In addition, in vitro experiments on mammalian AP-3 using liposomes or enriched Golgi membranes suggest Arf1 as an important factor in AP-3 recruitment, whereas acidic lipids do not have a major effect, in contrast to what was found for AP-1 and AP-2 (7, 11, 29, 30). Another study showed that membrane recruitment of AP-3 depends on the recognition of sorting signals in cargo tails and PI3P (31), similar to AP-1 recruitment via cargo tails, Arf1 and PI4P (32).However, since AP-1 and AP-3 are both recruited to the trans-Golgi network (TGN) in yeast (33), the mechanism of their recruitment likely differs. Even though Arf1 is required, yeast AP-3 seems to be present at the TGN before the arrival of the Arf1 guanine nucleotide exchange factor (GEF) Sec7 (33). This implies the necessity for additional factors at the TGN and a distinct mechanism to allow for spatial and temporal separation of AP-1 and AP-3 recruitment to membranes. Structural data on mammalian AP-1 and AP-2 “core” complexes without the hinge and ear domains of their large subunits revealed that both exist in at least two very defined conformational states: a “closed” cytosolic state, where the cargo-binding sites are buried within the complex, and an “open” state, where the same sites are available to bind cargo (7, 8, 10, 34, 35). Binding of Arf1 to AP-1 or PI(4,5)P₂ in case of AP-2 induces a conformational change in the complexes that enables them to bind cargo molecules carrying a conserved acidic di-Leucine or a Tyrosine-based motif, as for all three AP complexes in yeast (8, 34). Additional conformational states and intermediates have been reported for both, mammalian AP-1 and AP-2 complex. AP-1, for example, can be hijacked by the human immunodeficiency virus-1 (HIV-1) proteins viral protein u (Vpu) and negative factor (Nef), resulting in a hyper-open conformation of AP-1 (36, 37).An emerging model over the past years has suggested that APs have several binding sites that allow for the stabilization of membrane binding and the open conformation of the complexes, but there are initial interactions required that dictate their recruitment to the target membrane. Although these interaction sites for mammalian AP-1 and AP-2 have been identified in great detail based on interaction analyses and structural studies (8, 10, 11, 35, 36, 38, 39), structural data for AP-3 is largely missing. The C-terminal part of the μ-subunit of mammalian AP-3 has been crystallized together with a Yxxφ motif-containing a cargo peptide, which revealed a similar fold and cargo-binding site as shown for AP-1 and AP-2 (14). However, positively charged binding surfaces required for PIP-interaction were not well conserved. Although the “trunk” segment of AP-1 and AP-2 is known quite well by now, information on hinge and ear domains in context of these complexes is largely missing. Crystal structures of the isolated ear domains of α-, γ- and β2-adaptin have been published (40, 41, 42), and a study on mammalian AP-3 suggested a direct interaction between δ-ear and δ3 that interfered with Arf1-binding (43). Furthermore, during tethering of AP-3 vesicles with the yeast vacuole, the δ−subunit Apl5 of the yeast AP-3 complex binds to the Vps41 subunit of the HOPS complex as a prerequisite of fusion (18, 19, 21, 22).In this study, we applied single particle electron cryo-microscopy (cryo-EM) to analyze the purified full-length AP-3 complex from yeast and unraveled the factors required for AP-3 recruitment to membranes by biochemical reconstitution. Our data reveal that a surprisingly flexible AP-3 complex requires a combination of cargo, PI4P, and Arf1 for membrane binding, which explains its function in selective cargo sorting at the Golgi.     

Structural insights of mTOR complex 1

The mammalian target of rapamycin (mTOR), also known as the mechanistic target of rapamycin, is a central cell growth regulating kinase that forms large molecular complexes in all eukaryotic cells. A paper recently published in Science reports the architecture of mTOR complex 1 (mTORC1) and provides molecular insights into the regulation and substrate selectivity of mTORC1.The mammalian target of rapamycin (mTOR) exists in two different complexes, mTORC1 and mTORC2, which are distinguished by unique accessory protein Raptor and Rictor, respectively¹. Rapamycin is an mTORC1-specific inhibitor, which complexes with the FK506-binding 12 kDa protein (FKBP12) and inhibits Raptor-bound, but not Rictor-bound, mTOR. Rapamycin analogs have been used clinically to treat a number of human diseases, including cancer². A wide range of both extra- and intracellular signals, including growth factors, nutrient status and stress conditions, have been shown to regulate mTORC1 to control cell growth. Most notably, mTORC1 is hyperactivated by oncogenic PI3K-Akt signaling and promotes tumor growth¹.mTORC1 promotes cell growth through phosphorylation of a large number of cellular proteins, including the ribosomal S6 kinase 1 (S6K1) and eIF-4E-binding protein 1 (4E-BP1)³. Although mTORC2 shares the same catalytic kinase subunit with mTORC1, it phosphorylates substrates very different from those of mTORC1 and thus exerts different cellular functions. Despite the extensive studies, the mechanistic understanding of mTORC1 activation and substrate selectivity are rather limited, chiefly due to the lack of three dimensional structure of mTORC1. Understanding of mTORC1 molecular architecture is also of high importance for developing pharmacological drugs to target this pathway.Cryo-electron microscopy (cryo-EM) studies have shown that mTOR forms an obligate dimer with an overall rhomboid shape and a central cavity⁴. However, the reliability of the handedness of the reconstruction and the position of individual subunits was significantly compromised due to the low-resolution (26 Å) reconstruction. A subsequent study presented the 3.2 Å crystal structure of a complex of N-terminally truncated human mTOR and mLST8, which is a subunit commonly present in both mTORC1 and mTORC2. This crystal structure revealed more details of the structure of mTOR kinase domain as well as its inhibition by FKBP12-rapamycin complex⁵. However, information on the subunit arrangement within mTORC1 was missing because only a truncated mTOR fragment was analyzed and the Raptor subunit, which plays a key role in mTORC1 regulation and substrate selectivity, was lacking.In a recent paper published in Science, the architecture of human mTORC1 was revealed by high-resolution cryo-EM⁶. The authors purified mTORC1 complex (human mTOR together with Raptor, and mLST8) bound to FKBP12-rapamycin from insect cells. Single particle analysis of cryo-EM yielded a reconstruction with an overall resolution of 5.9 Å. To complement the reconstruction of mTORC1, the authors also resolved the structure of the fungus Chaetomium thermophilum Raptor (CtRaptor), which exhibits 44% sequence identity to human Raptor. The overall shape of the reconstruction agrees with that observed by low-resolution cyro-EM⁴; however, it appears that the handedness of the previous reconstruction was not assigned correctly. Generally, mTORC1 adopts a cage-like, dimeric architecture and appears in a hollow lozenge shape, in which Raptor and mLST8 contribute peripheral parts of the complex and make up the pinnacles of the longer and shorter axes of the lozenge, respectively. Interestingly, the N-terminus of mTOR, which was not resolved in previous study⁵, contains two α-helical solenoids. The larger section is a highly curved super-helix, which is named the “horn”, while the smaller region adopts a relatively linear arrangement and is referred to as the “bridge”⁶. Both sections are predominantly exposed to the environment, indicating a potential role in binding mTOR regulators. In addition, the horn and bridge HEAT domains pack against one another, and the first HEAT repeat of the horn region interlocks with the adjacent mTOR FAT domain, through which the two mTOR subunits forms a dimer independent of Raptor⁶. Another interesting observation is that the conformation of the kinase domain appears unaffected by dimerization, suggesting that the regulation of mTORC1 may be mainly through controlling substrate access to the active site.The authors further investigated how Raptor contributes to the formation of mTORC1 complex. Raptor interacts with mTOR through an α-solenoid stack formed between the horn and bridge domains of mTOR via the Raptor armadillo domain⁶. It is proposed that Raptor stabilizes the N-terminal region of mTOR by providing roughly two-thirds of the interaction surface with HEAT domains. As mentioned above, the formation of mTORC1 dimer is dependent on interaction of mTOR domains, but not Raptor, thus a model is suggested that Raptor binding may stabilize mTOR N-terminal conformation without directly engaging in dimer formation.The structure of mTORC1 also provides implications of mTORC1 substrate selectivity and delivery. Previous report has revealed that mTOR FRB domain and mLST8 prevent activity toward non-cognate substrates by limiting access to the ATP-binding cleft⁵. According to the current architecture, Raptor binding further restricts the access to the active site, resulting in the enclosure of the active site cleft from all directions and reduction of its width to ∼20 Å⁶. Moreover, binding of FKBP12-rapamycin complex to the FRB domain of mTORC1 further reduces the active site cleft to ∼10 Å⁶. Taken together, this model shows how architectural subunits of mTORC1 and FKBP12-rapamycin limit access to the recessed mTOR active site (Figure 1). It is notable that in contrary to previous findings⁴, this study indicates that FKBP12-rapamycin binding has no effect on mTORC1 stability.Open in a separate windowFigure 1Schematic model of mTORC1 substrate selectivity and delivery. (A) Substrate recruitment is dependent on their TOR signaling (TOS) motif binding to Raptor TOS-binding site. mTOR FRB domain and mLST8 prevent phosphorylation towards non-cognate substrates by limiting the access to the ATP-binding cleft. Raptor binding further restricts the access to the active site, resulting in the enclosure of the active site cleft from all directions. Only when the TOS motif present in the substrate is recognized by Raptor, substrates can be delivered to the mTOR kinase active site and phosphorylated. (B) The FKBP12-rapamycin complex binding to the FRB domain further blocks the active site cleft and prevents the access of substrate to kinase active site.In summary, the new structure reveals insights into the mTORC1 architecture and important clues for mTORC1 functional regulation. The kinase domain of mTOR maintains a constitutively active conformation at all times. Association with Raptor limits substrate accessibility to the mTOR kinase active site as the Raptor RNC domain is positioned directly at the mTOR active site cleft, thereby explaining how Raptor modulates substrate selectivity of mTORC1. Furthermore, the new structure explains mTORC1 inhibition by FKBP12-rapamycin through blocking substrate accessibility to the mTOR kinase active site. It should be noted that the architecture of mTORC1 still needs further improvement as the current resolution (5.9 Å) is not sufficient to reveal amino acid side chains of subunits and critical sites for dimer formation and activity control.     

Extracellular DNA Is Required for Root Tip Resistance to Fungal Infection

Plant defense involves a complex array of biochemical interactions, many of which occur in the extracellular environment. The apical 1- to 2-mm root tip housing apical and root cap meristems is resistant to infection by most pathogens, so growth and gravity sensing often proceed normally even when other sites on the root are invaded. The mechanism of this resistance is unknown but appears to involve a mucilaginous matrix or “slime” composed of proteins, polysaccharides, and detached living cells called “border cells.” Here, we report that extracellular DNA (exDNA) is a component of root cap slime and that exDNA degradation during inoculation by a fungal pathogen results in loss of root tip resistance to infection. Most root tips (>95%) escape infection even when immersed in inoculum from the root-rotting pathogen Nectria haematococca. By contrast, 100% of inoculated root tips treated with DNase I developed necrosis. Treatment with BAL31, an exonuclease that digests DNA more slowly than DNase I, also resulted in increased root tip infection, but the onset of infection was delayed. Control root tips or fungal spores treated with nuclease alone exhibited normal morphology and growth. Pea (Pisum sativum) root tips incubated with [³²P]dCTP during a 1-h period when no cell death occurs yielded root cap slime containing ³²P-labeled exDNA. Our results suggest that exDNA is a previously unrecognized component of plant defense, an observation that is in accordance with the recent discovery that exDNA from white blood cells plays a key role in the vertebrate immune response against microbial pathogens.Root diseases caused by soil-borne plant pathogens are a perennial source of crop loss worldwide (Bruehl, 1986; Curl and Truelove, 1986). These diseases are of increasing concern, as pesticides like methyl bromide are removed from the market due to environmental concerns (Gilreath et al., 2005). One possible alternative means of crop protection is to exploit natural mechanisms of root disease resistance (Nelson, 1990; Goswami and Punja, 2008; Shittu et al., 2009). Direct observation of root systems under diverse conditions has revealed that root tips, in general, are resistant to infection even when lesions are initiated elsewhere on the same plant root (Foster et al., 1983; Bruehl, 1986; Curl and Truelove, 1986; Smith et al., 1992; Gunawardena et al., 2005; Wen et al., 2007). This form of disease resistance is important for crop production because root growth and its directional movement in response to gravity, water, and other signals can proceed normally as long as the root tip is not invaded. The 1- to 2-mm apical region of roots houses the root meristems required for root growth and cap development, and when infection does occur, root development ceases irreversibly within a few hours even in the absence of severe necrosis (Gunawardena and Hawes, 2002). Mechanisms underlying root tip resistance to infection are unclear, but the phenomenon appears to involve root cap “slime,” a mucilaginous matrix produced by the root cap (Morré et al., 1967; Rougier et al., 1979; Foster, 1982; Chaboud, 1983; Guinel and McCully, 1986; Moody et al., 1988; Knee et al., 2001; Barlow, 2003; Iijima et al., 2008). Within the root cap slime of cereals, legumes, and most other crop species are specialized populations of living cells called root “border cells” (Supplemental Fig. S1; Hawes et al., 2000). Border cell numbers increase in response to pathogens and toxins such as aluminum, and the cell populations maintain a high rate of metabolic activity even after detachment from the root cap periphery (Brigham et al., 1995; Miyasaka and Hawes, 2000).As border cells detach from roots of cereals and legumes, a complex of more than 100 proteins, termed the root cap secretome, is synthesized and exported from living cells into the matrix ensheathing the root tip (Brigham et al., 1995). The profile of secreted proteins changes in response to challenge with soil-borne bacteria (De-la-Peña et al., 2008). In pea (Pisum sativum), root tip resistance to infection is abolished in response to proteolytic degradation of the root cap secretome (Wen et al., 2007). In addition to an array of antimicrobial enzymes and other proteins known to be components of the extracellular matrix and apoplast of higher plants, the DNA-binding protein histone H4 unexpectedly was found to be present among the secreted proteins (Wen et al., 2007). One explanation for the presence of histone is global leakage of material from disrupted nuclei in dead cells, but no cell death occurs during delivery of the secretome (Brigham et al., 1995; Wen et al., 2007). An alternative explanation for the presence of a secreted DNA-binding protein is that extracellular DNA (exDNA) also is present in root cap slime.exDNA has long been known to be a component of slimy biological matrices ranging from purulent localized human infections to bacterial capsules, biofilms, and snail exudate (Sherry and Goeller, 1950; Leuchtenberger and Schrader, 1952; Braun and Whallon, 1954; Smithies and Gibbons, 1955; Catlin, 1956; Fahy et al., 1993; Allesen-Holm et al., 2006; Spoering and Gilmore, 2006; Qin et al., 2007; Izano et al., 2008). Specialized white blood cells in humans and other species including fish recently have been shown to deploy a complex neutrophil extracellular “trap” (NET), composed of DNA and a collection of enzymes, in response to infection (Brinkmann et al., 2004; Brinkmann and Zychlinsky, 2007; Palić et al., 2007; Wartha et al., 2007; Yousefi et al., 2008). NETs appear to kill bacterial, fungal, and protozoan pathogens by localizing them within a matrix of antimicrobial peptides and proteins (Urban et al., 2006; Wartha et al., 2007; Guimaraes-Costa et al., 2009). Several extracellular peptides and proteins implicated in neutrophil function, including histone, also are present within the pea root cap secretome (Wen et al., 2007). exDNA linked with extracellular histone is a structural component of NETs, and treatment with DNase destroys NET integrity and function (Wartha et al., 2007). Moreover, human pathogens including group A Streptococcus and Streptococcus pneumoniae release extracellular DNase (Sherry and Goeller, 1950). When these activities are eliminated by mutagenesis of the encoding genes, bacteria lose their normal ability to escape the NET and multiply at the site of infection (Sumby et al., 2005; Buchanan et al., 2006). Here, we report that, in addition to histone and other secretome proteins, exDNA also is a component of root cap slime. When this exDNA is digested enzymatically, root tip resistance to infection is abolished.     

The K-Segment of Maize DHN1 Mediates Binding to Anionic Phospholipid Vesicles and Concomitant Structural Changes

Dehydrins (DHNs; late embryogenesis abundant D11 family) are a family of intrinsically unstructured plant proteins that accumulate in the late stages of seed development and in vegetative tissues subjected to water deficit, salinity, low temperature, or abscisic acid treatment. We demonstrated previously that maize (Zea mays) DHNs bind preferentially to anionic phospholipid vesicles; this binding is accompanied by an increase in α-helicity of the protein, and adoption of α-helicity can be induced by sodium dodecyl sulfate. All DHNs contain at least one “K-segment,” a lysine-rich 15-amino acid consensus sequence. The K-segment is predicted to form a class A2 amphipathic α-helix, a structural element known to interact with membranes and proteins. Here, three K-segment deletion proteins of maize DHN1 were produced. Lipid vesicle-binding assays revealed that the K-segment is required for binding to anionic phospholipid vesicles, and adoption of α-helicity of the K-segment accounts for most of the conformational change of DHNs upon binding to anionic phospholipid vesicles or sodium dodecyl sulfate. The adoption of structure may help stabilize cellular components, including membranes, under stress conditions.When plants encounter environmental stresses such as drought or low temperature, various responses take place to adapt to these conditions. Typical responses include increased expression of chaperones, signal transduction pathway and late embryogenesis abundant (LEA) proteins, osmotic adjustment, and induction of degradation and repair systems (Ingram and Bartels, 1996).Dehydrins (DHNs; LEA D11 family) are a subfamily of group 2 LEA proteins that accumulate to high levels during late stages of seed development and in vegetative tissues subjected to water deficit, salinity, low temperature, or abscisic acid (ABA) treatment (Svensson et al., 2002). Some DHNs are expressed constitutively during normal growth (Nylander et al., 2001; Rorat et al., 2004, 2006; Rodriguez et al., 2005). DHNs exist in a wide range of photosynthetic organisms, including angiosperms, gymnosperms, algae, and mosses (Svensson et al., 2002). DHNs are encoded by a dispersed multigene family and are differentially regulated, at least in higher plants. For example, 13 Dhn genes have been identified in barley (Hordeum vulgare), dispersed over seven genetic map locations (Choi et al., 1999; Svensson et al., 2002) and regulated variably by drought, low temperature, and embryo development (Tommasini et al., 2008). DHNs are localized in various subcellular compartments, including cytosol (Roberts et al., 1993), nucleus (Houde et al., 1995), chloroplast (Artus et al., 1996), vacuole (Heyen et al., 2002), and proximal to the plasma membrane and protein bodies (Asghar et al., 1994; Egerton-Warburton et al., 1997; Puhakainen et al., 2004). Elevated expression of Dhn genes generally has been correlated with the acquisition of tolerance to abiotic stresses such as drought (Whitsitt et al., 1997), salt (Godoy et al., 1994; Jayaprakash et al., 1998), chilling (Ismail et al., 1999a), or freezing (Houde et al., 1995; Danyluk et al., 1998; Fowler et al., 2001). The differences in expression and tissue location suggest that individual members of the Dhn multigene family have somewhat distinct biological functions (Close, 1997; Zhu et al., 2000; Nylander et al., 2001). Many studies have observed a positive correlation between the accumulation of DHNs and tolerance to abiotic stresses (Svensson et al., 2002). However, overexpression of a single DHN protein has not, in general, been sufficient to confer stress tolerance (Puhakainen et al., 2004).DHNs are subclassified by sequence motifs referred to as the K-segment (Lys-rich consensus sequence), the Y-segment (N-terminal conserved sequence), the S-segment (a tract of Ser residues), and the φ-segment (Close, 1996). Because of high hydrophilicity, high content of Gly (>20%), and the lack of a defined three-dimensional structure in the pure form (Lisse et al., 1996), DHNs have been categorized as “intrinsically disordered/unstructured proteins” or “hydrophilins” (Wright and Dyson, 1999; Garay-Arroyo et al., 2000; Tompa, 2005; Kovacs et al., 2008). On the basis of compositional and biophysical properties and their link to abiotic stresses, several functions of DHNs have been proposed, including ion sequestration (Roberts et al., 1993), water retention (McCubbin et al., 1985), and stabilization of membranes or proteins (Close, 1996, 1997). Observations from in vitro experiments include DHN binding to lipid vesicles (Koag et al., 2003; Kovacs et al., 2008) or metals (Svensson et al., 2000; Heyen et al., 2002; Kruger et al., 2002; Alsheikh et al., 2003; Hara et al., 2005), protection of membrane lipid against peroxidation (Hara et al., 2003), retention of hydration or ion sequestration (Bokor et al., 2005; Tompa et al., 2006), and chaperone activity against the heat-induced inactivation and aggregation of various proteins (Kovacs et al., 2008).Intrinsically disordered/unstructured proteins that lack a well-defined three-dimensional structure have recently been recognized to be prevalent in prokaryotes and eukaryotes (Oldfield et al., 2005). They fulfill important functions in signal transduction, gene expression, and binding to targets such as protein, RNA, ions, and membranes (Wright and Dyson, 1999; Tompa, 2002; Dyson and Wright, 2005). The disorder confers structural flexibility and malleability to adapt to changes in the protein environment, including water potential, pH, ionic strength, and temperature, and to undergo structural transition when complexed with ligands such as other proteins, DNA, RNA, or membranes (Prestrelski et al., 1993; Uversky, 2002). Structural changes from disorder to ordered functional structure also can be induced by the folding of a partner protein (Wright and Dyson, 1999; Tompa, 2002; Mouillon et al., 2008).The idea that DHNs interact with membranes is consistent with many immunolocalization studies, which have shown that DHNs accumulate near the plasma membrane or membrane-rich areas surrounding lipid and protein bodies (Asghar et al., 1994; Egerton-Warburton et al., 1997; Danyluk et al., 1998; Puhakainen et al., 2004). The K-segment is predicted to form a class A2 amphipathic α-helix, in which hydrophilic and hydrophobic residues are arranged on opposite faces (Close, 1996). The amphipathic α-helix is a structural element known to interact with membranes and proteins (Epand et al., 1995). Also, in the presence of helical inducers such as SDS and trifluoroethanol (Dalal and Pio, 2006), DHNs take on α-helicity (Lisse et al., 1996; Ismail et al., 1999b). We previously examined the binding of DHN1 to liposomes and found that DHNs bind preferentially to anionic phospholipids and that this binding is accompanied by an increase in α-helicity of the protein (Koag et al., 2003). Similarly, a mitochondrial LEA protein, one of the group III LEA proteins, recently has been shown to interact with and protect membranes subjected to desiccation, coupled with the adoption of amphipathic α-helices (Tolleter et al., 2007).Here, we explore the basis of DHN-vesicle interaction using K-segment deletion proteins. This study reveals that the K-segment is necessary and sufficient for binding to anionic phospholipid vesicles and that the adoption of α-helicity of DHN proteins can be attributed mainly to the K-segment.     

Maturation Stress Generation in Poplar Tension Wood Studied by Synchrotron Radiation Microdiffraction

Tension wood is widespread in the organs of woody plants. During its formation, it generates a large tensile mechanical stress, called maturation stress. Maturation stress performs essential biomechanical functions such as optimizing the mechanical resistance of the stem, performing adaptive movements, and ensuring long-term stability of growing plants. Although various hypotheses have recently been proposed, the mechanism generating maturation stress is not yet fully understood. In order to discriminate between these hypotheses, we investigated structural changes in cellulose microfibrils along sequences of xylem cell differentiation in tension and normal wood of poplar (Populus deltoides × Populus trichocarpa ‘I45-51’). Synchrotron radiation microdiffraction was used to measure the evolution of the angle and lattice spacing of crystalline cellulose associated with the deposition of successive cell wall layers. Profiles of normal and tension wood were very similar in early development stages corresponding to the formation of the S1 and the outer part of the S2 layer. The microfibril angle in the S2 layer was found to be lower in its inner part than in its outer part, especially in tension wood. In tension wood only, this decrease occurred together with an increase in cellulose lattice spacing, and this happened before the G-layer was visible. The relative increase in lattice spacing was found close to the usual value of maturation strains, strongly suggesting that microfibrils of this layer are put into tension and contribute to the generation of maturation stress.Wood cells are produced in the cambium at the periphery of the stem. The formation of the secondary wall occurs at the end of cell elongation by the deposition of successive layers made of cellulose microfibrils bounded by an amorphous polymeric matrix. Each layer has a specific chemical composition and is characterized by a particular orientation of the microfibrils relative to the cell axis (Mellerowicz and Sundberg, 2008). Microfibrils are made of crystalline cellulose and are by far the stiffest constituent of the cell wall. The microfibril angle (MFA) in each layer is determinant for cell wall architecture and wood mechanical properties.During the formation of wood cells, a mechanical stress of a large magnitude, known as “maturation stress” or “growth stress” (Archer, 1986; Fournier et al., 1991), occurs in the cell walls. This stress fulfills essential biomechanical functions for the tree. It compensates for the comparatively low compressive strength of wood and thus improves the stem resistance against bending loads. It also provides the tree with a motor system (Moulia et al., 2006), necessary to maintain the stem at a constant angle during growth (Alméras and Fournier, 2009) or to achieve adaptive reorientations. In angiosperms, a large tensile maturation stress is generated by a specialized tissue called “tension wood.” In poplar (Populus deltoides × Populus trichocarpa), as in most temperate tree species, tension wood fibers are characterized by the presence of a specific layer, called the G-layer (Jourez et al., 2001; Fang et al., 2008), where the matrix is almost devoid of lignin (Pilate et al., 2004) and the microfibrils are oriented parallel to the fiber axis (Fujita et al., 1974). This type of reaction cell is common in plant organs whose function involves the bending or contraction of axes, such as tendrils, twining vines (Bowling and Vaughn, 2009), or roots (Fisher, 2008).The mechanism at the origin of tensile maturation stress has been the subject of a lot of controversy and is still not fully understood. However, several recent publications have greatly improved our knowledge about the ultrastructure, chemical composition, molecular activity, mechanical state, and behavior of tension wood. Different models have been proposed and discussed to explain the origin of maturation stress (Boyd, 1972; Bamber, 1987, 2001; Okuyama et al., 1994, 1995; Yamamoto, 1998, 2004; Alméras et al., 2005, 2006; Bowling and Vaughn, 2008; Goswami et al., 2008; Mellerowicz et al., 2008). The specific organization of the G-layer suggests a tensile force induced in the microfibrils during the maturation process. Different hypotheses have been proposed to explain this mechanism, such as the contraction of amorphous zones within the cellulose microfibrils (Yamamoto, 2004), the action of xyloglucans during the formation of microfibril aggregates (Nishikubo et al., 2007; Mellerowicz et al., 2008), and the effect of changes in moisture content stimulated by pectin-like substances (Bowling and Vaughn, 2008). A recent work (Goswami et al., 2008) argued an alternative model, initially proposed by Münch (1938), which proposed that the maturation stress originates in the swelling of the G-layer during cell maturation and is transmitted to the adjacent secondary layers, where the larger MFAs allow an efficient conversion of lateral stress into axial tensile stress. Although the proposed mechanism is not consistent with the known hygroscopic behavior of tension wood, which shrinks when it dries and not when it takes up water (Clair and Thibaut, 2001; Fang et al., 2007; Clair et al., 2008), this hypothesis focused attention on the possible role of cell wall layers other than the G-layer. As a matter of fact, many types of wood fibers lacking a G-layer are known to produce axial tensile stress, such as normal wood of angiosperms and conifers (Archer, 1986) and the tension wood of many tropical species (Onaka, 1949; Clair et al., 2006b; Ruelle et al., 2007), so that mechanisms strictly based on an action of the G-layer cannot provide a general explanation for the origin of tensile maturation stress in wood.In order to further understanding, direct observations of the mechanical state of the different cell wall layers and their evolution during the formation of the tension wood fibers are needed. X-ray diffraction can be used to investigate the orientation of microfibrils (Cave, 1966, 1997a, 1997b; Peura et al., 2007, 2008a, 2008b) and the lattice spacing of crystalline cellulose. The axial lattice spacing d₀₀₄ is the distance between successive monomers along a cellulose microfibril and reflects its state of mechanical stress (Clair et al., 2006a; Peura et al., 2007). If cellulose microfibrils indeed support a tensile stress, they should be found in an extended state of deformation. Under this assumption, the progressive development of maturation stress during the cell wall formation should be accompanied by an increase in cellulose lattice spacing. Synchrotron radiation allows a reduction in the size of the x-ray beam to some micrometers while retaining a strong signal, whereby diffraction analysis can be performed at a very local scale (Riekel, 2000). This technique has been used to study sequences of wood cell development (Hori et al., 2000; Müller et al., 2002). In this study, we report an experiment where a microbeam was used to analyze the structural changes of cellulose in the cell wall layers of tension wood and normal wood fibers along the sequence of xylem cell differentiation extending from the cambium to mature wood (Fig. 1). The experiment was designed to make this measurement in planta, in order to minimize sources of mechanical disturbance and be as close as possible to the native mechanical state (Clair et al., 2006a). The 200 and 004 diffraction patterns of cellulose were analyzed to investigate the process of maturation stress generation in tension wood.Open in a separate windowFigure 1.Schematic of the experimental setup, showing the x-ray beam passing perpendicular to the longitudinal-radial plane of wood and the contribution of the 004 and 200 crystal planes to the diffraction pattern recorded by the camera. [See online article for color version of this figure.]     

ATG8ylation of proteins: A way to cope with cell stress?

The ATG8 family of proteins regulates autophagy in a variety of ways. Recently, ATG8s were demonstrated to conjugate directly to cellular proteins in a process termed “ATG8ylation,” which is amplified by mitochondrial damage and antagonized by ATG4 proteases. ATG8s may have an emerging role as small protein modifiers.

ATG8 proteins directly conjugate to cellular proteinsAutophagy describes the capture of intracellular material by autophagosomes and their delivery to lysosomes for destruction (Kaur and Debnath, 2015). This process homeostatically remodels the intracellular environment and is necessary for an organism to overcome starvation (Kaur and Debnath, 2015). The autophagy pathway is coordinated by autophagy-related (ATG) proteins that are controlled by diverse post-translational modifications (e.g., phosphorylation, acetylation, ubiquitination, and lipidation; Ichimura et al., 2000; McEwan and Dikic, 2011). Recently, a previously uncharacterized post-translational modification termed “ATG8ylation” was uncovered (Agrotis et al., 2019; Nguyen et al., 2021). ATG8ylation is the direct covalent attachment of the small ubiquitin-like family of ATG8 proteins to cellular proteins (Agrotis et al., 2019; Nguyen et al., 2021). Until now, the only known instances of ATG8 conjugation to proteins were of a transient nature, as E1- and E2-like intermediates with ATG7 and ATG3, respectively, as a way of ligating ATG8 to the lipid phosphatidylethanolamine during autophagy (Ichimura et al., 2000). Therefore, ATG8ylation may represent an underappreciated regulatory mechanism for many cellular proteins that coordinate pathways such as mitophagy.ATG8s play many roles in the autophagy pathwayDuring canonical autophagy, the ATG8 family (comprising LC3A, -B, and -C and GABARAP, -L1, and -L2) undergoes molecular processing that concludes with their attachment to phosphatidylethanolamine, enabling proper construction of autophagosomes and subsequent autophagosome–lysosome fusion (Nguyen et al., 2016). The ATG4 family of cysteine proteases (ATG4A, -B, -C, and -D) cleaves ATG8 proteins immediately after a conserved glycine residue in their C terminus in a process dubbed “priming,” which leads to the formation of ATG8-I (Skytte Rasmussen et al., 2017; Tanida et al., 2004). ATG7 then attaches to the exposed glycine residue of ATG8-I via a thioester linkage to form an E1 ubiquitin-like complex that transfers ATG8-I to ATG3 in a similar way to generate an E2-like complex (Ichimura et al., 2000). The ATG5–ATG12–ATG16L1 complex then catalyzes the E3-like transfer of ATG8-I from ATG3 to phosphatidylethanolamine to form ATG8-II, which is the lipidated species that is incorporated into double membrane–bound compartments such as autophagosomes (Hanada et al., 2007). The lipidation of ATG8s and their recruitment to the phagophore are not essential for the formation of autophagosomes but are important for phagophore expansion, the selective capture of autophagic substrates, and autophagosome–lysosome fusion (Kirkin and Rogov, 2019; Nguyen et al., 2016). Intriguingly, ATG8 lipidation is multifaceted, as ATG8s can be alternatively lipidated with phosphatidylserine (instead of phosphatidylethanolamine) to enable their recruitment to single membrane–bound compartments during LC3-associated phagocytosis, influenza infection, and lysosomal dysfunction (Durgan et al., 2021).The discovery of ATG8ylationKey insights into ATG8ylation came from the observation that various ATG8s form high-molecular-weight species in cells following the expression of their primed forms that have their C-terminal glycine exposed (for example, LC3B-G), bypassing the need for cleavage by ATG4 (Agrotis et al., 2019; Nguyen et al., 2021). Indeed, on an immunoblot, ATG8⁺ “smears” resemble that of ubiquitinated proteins (Agrotis et al., 2019; Nguyen et al., 2021). Traditionally, in the autophagy field, ATG8⁺ smears were thought to arise from poor antibody specificity. However, in light of recent findings, this widely accepted interpretation has been challenged, given that ATG8⁺ smears are enriched following ATG8 overexpression and disappear in the absence of ATG8s (Agrotis et al., 2019; Nguyen et al., 2021). Smearing has also been detected after immunoprecipitation of epitope-tagged ATG8s from cell extracts under denaturing conditions, ruling out noncovalent interactions accounting for this upshift (Agrotis et al., 2019; Nguyen et al., 2021). Further, smearing is not abolished by deubiquitinase treatment, arguing strongly against ATG8 ubiquitination as the cause (Nguyen et al., 2021). Everything considered, the most plausible explanation is that ATG8 itself undergoes covalent linkage to cellular proteins, akin to ubiquitin and NEDD8 modifiers, which are structurally similar to ATG8s. Remarkably, the protease ATG4 antagonizes the ATG8ylation state of many proteins (Agrotis et al., 2019; Nguyen et al., 2021).ATG4 displays isoform-specific proteolytic cleavage of ATG8ATG4 is required for the formation of autophagosomes, but its protease activity is not (Nguyen et al., 2021). The protease activity of ATG4 is, however, required for ATG8 processing, such as priming ahead of lipidation and de-lipidation, which removes excess ATG8 from autophagosomes and other membranes (Nguyen et al., 2021; Tanida et al., 2004; Fig. 1 A). Apart from these functions, ATG4 regulates the deubiquitinase-like removal of ATG8 from cellular proteins (de-ATG8ylation; Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 A). Consistent with this role, deletion of all four ATG4 isoforms (A, B, C, and D) increases the abundance of ATG8ylated proteins (Nguyen et al., 2021). In contrast, overexpression of ATG4B has the opposite effect, but only if its protease activity is intact (Agrotis et al., 2019). As such, ATG4 inhibits the ATG8ylation state of many proteins, which is likely to modulate their downstream functions.Open in a separate windowFigure 1.The many roles of ATG4 in ATG8 processing. (A) Molecular processing of ATG8 proteins by ATG4 illustrating its roles in priming, de-lipidation, and de-ATG8ylation. The structure of LC3B (Protein Data Bank accession no. 1V49) was used to denote ATG8 (G, glycine; PE, phosphatidylethanolamine). (B) Heatmap summarizing relationships between ATG4 isoforms and ATG8 family members. Data were summarized for qualitative interpretation (Agrotis et al., 2019; Li et al., 2011; Nguyen et al., 2021). Int., intermediate; N.d., not determined. (C) Graphical summary of questions moving forward with ATG8ylation (P, phosphorylation).ATG4 is an important “gatekeeper” for ATG8 conjugation events. ATG4 primes ATG8s to expose their C-terminal glycine, which is required for conjugation to proteins or lipids; however, ATG4 also catalyzes de-ATG8ylation and de-lipidation events, respectively (Agrotis et al., 2019; Nguyen et al., 2021; Tanida et al., 2004). Because the C-terminal glycine of a single ATG8 is occupied when conjugated to a protein or lipid, it is unlikely that ATG8ylated proteins directly engage with phagophore membranes in the same way as ATG8-II. Indeed, protease protection assays with recombinant ATG4B reveal that de-ATG8ylation of cell lysates remains unchanged with or without organellar membrane disruption, suggesting that ATG8ylated proteins are largely cytoplasmic facing rather than intraluminal (Agrotis et al., 2019). Paradoxically, however, ATG8ylation is enhanced by lysosomal V-type ATPase inhibition, which blocks the degradation of lysosomal contents, indicating that ATG8ylated substrates may undergo lysosome-dependent turnover (Agrotis et al., 2019; Nguyen et al., 2021). One explanation for these differences may be that the process of ATG8ylation is itself sensitive to lysosomal dysfunction.Functional relationships between ATG4s and ATG8sIsoforms of ATG4 show clear preferences for proteolytically processing ATG8 subfamilies (i.e., LC3s and GABARAPs) for de-ATG8ylation and priming upstream of phosphatidylethanolamine ligation (Agrotis et al., 2019; Li et al., 2011; Nguyen et al., 2021; Fig. 1 B). ATG4A strongly reduces the abundance of proteins that have been ATG8ylated with the GABARAP family while promoting ligation of GABARAPs to phosphatidylethanolamine (Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 B). In contrast, ATG4B strongly reduces the abundance of proteins that have been ATG8ylated with LC3 proteins while promoting ligation of LC3s to phosphatidylethanolamine (Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 B). In comparison, ATG4C and -D lack obvious de-ATG8ylation activity, although the latter weakly promotes phosphatidylethanolamine ligation to GABARAPL1 only (Nguyen et al., 2021). These functional similarities between ATG4 isoforms are consistent with both their sequence and structural homology (i.e., ATG4A and -B are most similar; Maruyama and Noda, 2018; Satoo et al., 2009). Structurally, ATG4B adopts an auto-inhibited conformation with its regulatory loop and N-terminal tail blocking substrate entry to its proteolytic core (Maruyama and Noda, 2018). LC3B induces conformational rearrangements in ATG4B that involve displacement of its regulatory loop and its N-terminal tail, with the latter achieved by an interaction between the ATG8-interacting region in its N-terminal tail with a second copy of LC3B that functions allosterically (Maruyama and Noda, 2018; Satoo et al., 2009). These rearrangements permit entry of LC3B into the proteolytic core of ATG4B, where cleavage of LC3B following its C-terminal glycine occurs (Li et al., 2011; Maruyama and Noda, 2018). ATG4B^L232 is directly involved in LC3B binding and its selectivity for LC3s (Satoo et al., 2009). This residue corresponds to ATG4A^I233 and, when substituted for leucine, gives ATG4A^I233L the ability to efficiently process LC3 proteins, whereas without this mutation it preferentially processes GABARAPs (Satoo et al., 2009). Moreover, the ATG8–ATG4 interaction is necessary for the de-ATG8ylation of cellular proteins, as an LC3B-G^Q116P mutant that cannot bind to ATG4 leads to widespread ATG8ylation (Agrotis et al., 2019). Altogether, these observations hint toward a common mechanism of ATG8 cleavage that regulates priming, de-lipidation, and de-ATG8ylation.Mitochondrial damage promotes ATG8ylationATG8ylation of cellular proteins appears to be enhanced by mitochondrial depolarization and inhibition of the lysosomal V-type ATPase (Agrotis et al., 2019; Nguyen et al., 2021). This may be the consequence of acute ATG4A and -B inhibition, given that cells lacking all ATG4 isoforms display an increased abundance of ATG8ylated proteins and are insensitive to further increase by mitochondrial depolarization or lysosomal V-type ATPase inhibition (Agrotis et al., 2019; Nguyen et al., 2021). Indeed, mitochondrial depolarization leads to activation of ULK1, which phosphorylates ATG4B^S316 to inhibit its protease activity (Pengo et al., 2017). Similarly, mitochondrial depolarization stimulates TBK1 activation, which prevents de-lipidation of ATG8s by blocking the ATG8–ATG4 interaction through phosphorylation of LC3C^S93/S96 and GABARAP-L2^S87/S88 (Herhaus et al., 2020; Richter et al., 2016). As such, ATG8 phosphorylation may render ATG8ylated substrates more resistant to de-ATG8ylation by ATG4s. This may be analogous to how chains of phosphorylated ubiquitin^S65 are more resistant to hydrolysis by deubiquitinating enzymes than unphosphorylated ones (Wauer et al., 2015). Moreover, ATG8ylation is insensitive to nutrient deprivation and pharmacological inhibition of mTOR, which rules out a functional contribution of this process to starvation-induced autophagy (Agrotis et al., 2019). Therefore, ATG8ylation may be a unique aspect of mitophagy (and perhaps also other forms of selective autophagy) given that depolarization potently activates Parkin-dependent mitophagy (Agrotis et al., 2019; Nguyen et al., 2021).Substrates of ATG8ylationBased on ATG8⁺ smearing, ATG4 regulates the de-ATG8ylation of numerous proteins (Agrotis et al., 2019; Nguyen et al., 2021). For the majority, their identity, induced structural and functional changes, and the cellular contexts during which these modifications occur await exploration. Considering that the ATG8 interactome is well characterized, it is likely that at least some ATG8ylated proteins have been mistaken for ATG8-binding partners (Behrends et al., 2010). Given their E2- and E3-like roles in ATG8 lipidation, it is remarkable that ATG3 and ATG16L1 are themselves modified by ATG8ylation (Agrotis et al., 2019; Hanada et al., 2007; Ichimura et al., 2000; Nguyen et al., 2021). Lysine mutagenesis indicates that ATG3^K243 is the “acceptor” site for ATG8ylation (Agrotis et al., 2019). ATG3^K243 is essential for its conjugation to either LC3B or ATG12 and is required for autophagosomes to form around damaged mitochondria (Agrotis et al., 2019; Radoshevich et al., 2010). This also raises the possibility that key functions originally attributed to ATG3–ATG12 conjugation may be, at least in part, due to ATG3–ATG8 conjugation. Because multiple high-molecular-weight species of ATG3 are enriched following immunoprecipitation of primed LC3B-G from cells lacking ATG4B, it is likely that ATG3 is either mono-ATG8ylated at several sites or poly-ATG8ylated (Agrotis et al., 2019). ATG8ylation of ATG3 may also reflect the stabilization of its E2-like intermediate (Ichimura et al., 2000). ATG8ylation of ATG16L1 may regulate whether canonical or noncanonical autophagy pathways are activated (Durgan et al., 2021; Nguyen et al., 2021). In line with this possibility, the WD40 domain mutant of ATG16L1^K490A prevents lipidation of ATG8s with phosphatidylserine (i.e., during noncanonical autophagy pathways) but not phosphatidylethanolamine (i.e., during canonical autophagy; Durgan et al., 2021). Moreover, given that ATG8ylation of protein targets correlates with the activation of mitophagy, it is tempting to speculate that it may stimulate the E2-/E3-like activity of the ATG8 conjugation machinery to amplify mitochondrial capture and destruction.Concluding remarksThe finding that numerous cellular proteins are modified by ATG8ylation poses several questions about how signaling networks are coordinated during selective autophagy (i.e., mitophagy). Whether ATG8ylation is augmented by mitochondrial injury per se or is the consequence of mitophagy activation is yet to be determined, as is whether this phenomenon occurs during other types of selective autophagy (e.g., ER-phagy, ribophagy, and lysophagy; Kirkin and Rogov, 2019; Fig. 1 C). While the in vivo relevance of ATG8ylation is not yet understood, it is plausible that this process could be altered in diseases with defective mitophagy (e.g., Parkinson’s disease and atherosclerosis). Exploring the mechanistic aspects of ATG8ylation (e.g., ATG8 ligases and regulatory proteins, linkage types, acceptor sites, etc.) and de-ATG8ylation by ATG4 will improve our understanding about how this modifier alters the structure and biological function of cellular proteins (Fig. 1 C). By identifying ATG8ylated substrates, or the ATG8ylome, insights into whether ATG8ylation is a ubiquitous epiphenomenon or a post-translational modification that is selective to proteins of distinct biological function(s) will become clearer (Fig. 1 C). Considering the similarity of ATG8s with bona fide modifier proteins (e.g., ubiquitin and ubiquitin-like proteins) and the diversity of their substrates (e.g., lipid species and proteins), only now are we beginning to understand the functional complexities of the ATG8 protein family.     

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

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

MRAP2 inhibits β-arrestin recruitment to the ghrelin receptor by preventing GHSR1a phosphorylation

The melanocortin receptor accessory protein 2 (MRAP2) is essential for several physiological functions of the ghrelin receptor growth hormone secretagogue receptor 1a (GHSR1a), including increasing appetite and suppressing insulin secretion. In the absence of MRAP2, GHSR1a displays high constitutive activity and a weak G-protein–mediated response to ghrelin and readily recruits β-arrestin. In the presence of MRAP2, however, G-protein–mediated signaling via GHSR1a is strongly dependent on ghrelin stimulation and the recruitment of β-arrestin is significantly diminished. To better understand how MRAP2 modifies GHSR1a signaling, here we investigated the role of several phosphorylation sites within the C-terminal tail and third intracellular loop of GHSR1a, as well as the mechanism behind MRAP2-mediated inhibition of β-arrestin recruitment. We show that Ser²⁵² and Thr²⁶¹ in the third intracellular loop of GHSR1a contribute to β-arrestin recruitment, whereas the C-terminal region is not essential for β-arrestin interaction. Additionally, we found that MRAP2 inhibits GHSR1a phosphorylation by blocking the interaction of GRK2 and PKC with the receptor. Taken together, these data suggest that MRAP2 alters GHSR1a signaling by directly impacting the phosphorylation state of the receptor and that the C-terminal tail of GHSR1a prevents rather than contribute to β-arrestin recruitment.

The “hunger hormone” ghrelin is secreted by X/A cells of the oxyntic mucosa of the stomach in response to a low energetic state, which leads to an increase in appetite (1, 2) and prevents hypoglycemia (3, 4). Ghrelin is the agonist of the growth hormone secretagogue receptor 1a (GHSR1a), a G-protein–coupled receptor (GPCR) expressed in the brain and in multiple peripheral organs including the heart and the endocrine pancreas. Activation of GHSR1a by ghrelin in hypothalamic agouti-related protein (AgRP) neurons potently stimulates feeding (5, 6, 7). In pituitary somatotrophs, GHSR1a stimulation promotes growth hormone release (8, 9, 10). Finally, in cardiomyocytes, ghrelin increases cell survival and contractility (11, 12) while in the endocrine pancreas the hormone inhibits insulin secretion (13, 14).GHSR1a primarily couples to Gα_q/11, thus stimulating the production of intracellular inositol triphosphate (IP) 3. Like other GPCRs, agonist stimulation results in phosphorylation of GHSR1a by kinases, including GPCR kinase 2 (GRK2) and PKC (15), and β-arrestin recruitment. Notably, GHSR1a contains several phosphorylation sites within the C-terminal tail, some of which have been shown to be important for β-arrestin recruitment (16). However, although other putative phosphorylation sites are present in the third intracellular loop (ICL3) of GHSR1a, their role in β-arrestin recruitment has not yet been described.When expressed in heterologous cells, GHSR1a displays a high constitutive activity and a limited ghrelin-stimulated responses (17).Both constitutive- and agonist-stimulated GHSR1a signaling are regulated by the single transmembrane melanocortin receptor accessory protein 2 (MRAP2), which functions to drastically reduce GHSR1a constitutive activity and increase ghrelin-stimulated responses (17). Additionally, MRAP2 significantly inhibits ghrelin-induced β-arrestin recruitment to GHSR1a (17). As such, MRAP2 is essential for several physiological functions of ghrelin including its orexigenic activity (18) and its insulinostatic actions (14). Global or AGRP neuron–targeted deletion of MRAP2 abrogates the effect of ghrelin on food intake (18) and global or pancreatic δ-cell-targeted deletion of MRAP2 prevents ghrelin-mediated inhibition of insulin secretion (14).Although expressed in AGRP neurons and pancreatic δ-cells (thus promoting G-protein coupling and inhibiting β-arrestin-dependent signaling), MRAP2 is not present in every GHSR1a-expressing tissue. Consequently, it is possible that β-arrestin signaling plays an important role in the physiological function of ghrelin in tissues where MRAP2 is absent. Whereas, the inhibition of β-arrestin recruitment to GHSR1a by MRAP2 is well established and the domains of MRAP2 required for this function have been identified (17), the molecular mechanism by which MRAP2 alters GHSR1a signaling is not yet understood. In this study, we investigated the importance of GHSR1a phosphorylation for β-arrestin recruitment and the mechanism involved in MRAP2-mediated inhibition of β-arrestin recruitment.     

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

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

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