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.     

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.     

Louis Reichardt: The long climb to science's summits

From the highest mountains to biology''s own Everest—the brain—Reichardt tackles the biggest challenges of climbing and biology.Louis Reichardt''s scientific career has spanned the simple and the complex. As a graduate student, Reichardt helped to uncover the now renowned DNA regulatory mechanisms that allow one of the simplest life forms, lambda phage, either to hide within a cell or to make its presence known via massive replication (1).Open in a separate windowLouis ReichardtLetting his curiosity for the unknown guide him, Reichardt then forayed into a much more intricate system, the brain. As a postdoc, he showed that growth conditions influenced which neurotransmitters are synthesized by isolated neurons (2). Later, as a professor at UCSF, he discovered synaptotagmin, using the first monoclonal antibody that defined a synaptic vesicle membrane protein (3), showed that expression levels of nerve growth factor in target tissues correlate with the density of innervation (4), and characterized the properties of mice lacking genes encoding the neurotrophins and their Trk receptors (5, 6).Now a professor and director of the Neuroscience Program at UCSF, Reichardt''s laboratory still studies the interface of cell biology and neurobiology, including the involvement of cell adhesion molecules in synaptic development (7). He explains that science is not so different from his favorite hobby, mountain climbing.

“The gist of it was ‘Others more foolish might try this, but it was not for us.’ I thought, ‘We''ll obviously have to do it.’”

     

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.     

In-cell infection: bringing uninvited guests

Cell-in-cell structures resulting from live cell engulfment were identified more than 100 years ago, but their physiological significance has remained largely obscure. Now Ni et al. identify a new role for cell-in-cell structure formation, called “in-cell infection” that spreads Epstein-Barr virus from infected B cells to epithelial cells, an activity that may predispose to cancer.Epstein-Barr virus (EBV) is a common herpesvirus infecting up to 90% or more of the human population that causes mononucleosis, is associated with autoimmune conditions, and predisposes to cancer¹. EBV persists as a latent infection within B cells and predisposes to cancers of B cell origin, including Hodgkin''s and Burkett''s lymphoma, due to expression of latency genes, which leads to B cell transformation. Infected individuals are also predisposed to developing nasopharyngeal and gastric carcinoma, as epithelial cells also harbor latent EBV. However, while the mechanism of EBV entry into B cells is well characterized, how EBV infects epithelium has remained obscure. In a recent paper published in Cell Research, Ni et al.² identify a novel mechanism for EBV infection of epithelial cells, which they term “in-cell infection”, an insidious mode of viral entry that takes advantage of whole cell ingestion.Viral infection is generally mediated by viral envelope glycoproteins that bind to specific receptors on target cells, leading to membrane fusion and viral entry. To infect B cells, the EBV envelope protein gp350 binds to the complement receptor 2 (CR2) on target cells, followed by interaction of gp42 with MHC class II molecules, and virus-to-target cell fusion is mediated by gp42, gH and gL proteins³. Unlike B cells, epithelial cells do not normally express complement receptors or MHC class II molecules, and are generally not infected by purified EBV. Previously described alternative modes of EBV infection of epithelial cells include “cell-to-cell” and “transfer” infection, where B cells have been found to act as carriers to mediate infection through cell adhesion protein-dependent conjugation^3,4,5,6.Ni et al.² now describe a different mode of epithelial cell infection, called “in-cell infection”, that occurs by ingestion of whole EBV-infected B cells, leading to the formation of “cell-in-cell” structures. B cell ingestion in this context resembles “entosis”, a mechanism previously found to mediate cell-in-cell structure formation in epithelial cultures and human tumors⁷. Entosis also promotes the uptake of hematopoietic cells into epithelial cells or cancer cells of various types^8,9. Incredibly, the authors find that entosis-like internalization of latent EBV-infected B cells (Akata) into cultured nasopharyngeal carcinoma cells (CNE-2) leads to the activation of EBV and the transfer of virus to host (CNE-2) cells. Cells infected in this manner express viral gene products, and produce virions upon stimulation that can infect naïve cells of either B cell or epithelial cell origin, indicating potent infection ability and altered tropism of EBV produced by this mechanism.Frequent cell-in-cell structure formation involving EBV-infected B cells is shown by the authors to occur in clinical nasopharyngeal carcinoma samples, suggesting that the in-cell infection mechanism is a likely contributor to viral spread in vivo, and may be linked to carcinoma development. Intriguingly, entosis itself may participate in tumorigenesis by promoting aneuploidy¹⁰, and by supplying cancer cells with nutrients¹¹. As the authors found that EBV infection promoted entosis-like cell uptake, this mode of viral spread could affect tumorigenesis by multiple mechanisms. For in-cell infection, it seems that the nutrients taken in upon the death of internalized cells come mixed with virus that is insidiously transferred to hosts, in a manner perhaps like a Trojan horse enterring with a hidden viral payload (Figure 1).Open in a separate windowFigure 1In-cell infection delivers virus to insusceptible host cells. The B cell infected by EBV resembles a Trojan horse that delivers a hidden viral payload to host epithelial cells.The identification of in-cell infection by Ni et al.² makes a significant contribution to cell-in-cell research by identifying a new pathophysiological role for an entosis-like process. Cell-in-cell structures were first reported over 100 years ago, but the mechanisms that control the formation of such structures and their significance are only now starting to emerge¹². As is often the case with groundbreaking research, the discovery of in-cell infection² raises many new interesting questions. What is the mechanism of B cell internalization into epithelial cells? Entosis is previously described to involve cell adhesion receptors, such as E-cadherin, and Rho-kinase that promotes the actomyosin contraction that drives cell uptake⁷. The molecular mechanism controlling the entry of EBV-infected B cells into epithelial cells will be important to uncover, as other mechanisms in addition to entosis can also mediate the uptake of live cells¹². How is EBV activated by the formation of cell-in-cell structures? How is EBV transferred from internalized cells to hosts? And importantly, can other viruses, such as HIV, spread by in-cell infection? The answers to these questions await further research.     

An emerging consensus on voltage-dependent gating from computational modeling and molecular dynamics simulations

Developing an understanding of the mechanism of voltage-gated ion channels in molecular terms requires knowledge of the structure of the active and resting conformations. Although the active-state conformation is known from x-ray structures, an atomic resolution structure of a voltage-dependent ion channel in the resting state is not currently available. This has motivated various efforts at using computational modeling methods and molecular dynamics (MD) simulations to provide the missing information. A comparison of recent computational results reveals an emerging consensus on voltage-dependent gating from computational modeling and MD simulations. This progress is highlighted in the broad context of preexisting work about voltage-gated channels.Voltage-gated K⁺ (K_V) channels and prokaryotic voltage-gated Na⁺ (Na_V) channels are formed by four subunits surrounding a central aqueous pore that allows ion permeation. Each subunit consists of six transmembrane α-helical segments called S1 to S6; the first four of these, S1–S4, constitute the voltage-sensor domain (VSD), whereas the S5–S6 segments assemble to form an ion-selective pore domain (see Fig. 1). The VSDs respond to changes in the potential difference across the cell membrane. When the membrane is depolarized, the VSD in each subunit undergoes a conformational transition from a resting to an activated state, and this information is communicated to the ion-conducting pore to promote its opening (Bezanilla et al., 1994; Zagotta et al., 1994). The activation of the VSD and opening of the pore are associated with the transfer of an electric charge ΔQ across the membrane, called the “gating charge” (Sigworth, 1994). Opening of the voltage-gated K⁺ channel Shaker corresponds to the outward translocation of a large positive charge on the order of 12–14 elementary charges (Schoppa et al., 1992). Four highly conserved arginines along S4 (R1, R2, R3, and R4) underlie the dominant contributions to the total gating charge of Shaker and appear to be mainly responsible for the coupling to the membrane voltage (Papazian et al., 1991; Aggarwal and MacKinnon, 1996; Seoh et al., 1996). The overall structure of eukaryotic voltage-gated Na⁺ channels, which are composed of four analogous subunits covalently linked in a single polypeptide, appears to be similar (Catterall, 2012).Open in a separate windowFigure 1.Overall view of the voltage-activated Kv1.2 K⁺ channel (Protein Data Bank accession no. 3LUT). (A) Two of the four subunits of the channel are displayed from a side view. The VSD comprises the transmembrane segments S1–S4, and the pore domain comprises the transmembrane segments S5–S6. (B) The tetramer is displayed from the extracellular side (each subunit is a different color). The two views are related by a 90° rotation.The nature of the conformational change within the VSD, and how it is communicated to the pore domain, is the key question that must be answered to explain voltage-dependent gating. Ultimately, we need to know the 3-D structure of the multiple resting and activated states of the VSDs and their relationship to the closed and open conformations of the pore at atomic resolution to understand the voltage-dependent gating mechanism in molecular terms. However, although x-ray crystallographic structures of the Kv1.2 channel, Kv1.2/Kv2.1 chimera, and bacterial Na_VAb channels have provided information on the conformation of the active state (Long et al., 2005, 2007; Payandeh et al., 2011), no atomic resolution structure of a K_V or Na_V channel in the resting state is currently available. This has motivated the use of computations to provide the missing information about channel gating (Yarov-Yarovoy et al., 2006, 2012; Pathak et al., 2007; Bjelkmar et al., 2009; Delemotte et al., 2010, 2011; Khalili-Araghi et al., 2010, 2012; Schwaiger et al., 2011; Vargas et al., 2011; Jensen et al., 2012). These computational studies have relied on different approaches, including Rosetta modeling, a protein-folding method using knowledge-based potentials, and molecular dynamics (MD) simulations, consisting of propagating Newton’s classical equation of motion as a function of time using an all-atom force field. Remarkably, despite the considerable variations in computational methodologies and in template x-ray structures used, a highly consistent picture is emerging from these studies. Here we briefly review the most recent results in the broad context of preexisting work about voltage-gated channels.

Computational models of the resting state

Early models of the resting-state conformation of the VSD were obtained using the Rosetta method (Yarov-Yarovoy et al., 2006; Pathak et al., 2007); these initial models were subsequently refined with all-atom MD simulations (Khalili-Araghi et al., 2010) and with high-resolution Rosetta algorithms (Yarov-Yarovoy et al., 2012). Independent studies obtained very similar conformations using a combination of experimentally derived constraints based on engineered cross-links and metal bridges during MD simulations (Delemotte et al., 2010, 2011; Henrion et al., 2012). Moreover, these earliest models (Yarov-Yarovoy et al., 2006; Pathak et al., 2007) predicted pairs of neighboring residues before they were identified experimentally (Campos et al., 2007). Subsequent refinement of structural models of the resting state made it possible to demonstrate the existence of a consensus 3-D conformation of the VSD that satisfied a wide range of experimental data (Vargas et al., 2011). Rosetta models for the bacterial sodium channel NaChBac are very similar to those of K_V channels and have been extensively tested by disulfide cross-linking studies (DeCaen et al., 2008, 2009, 2011; Yarov-Yarovoy et al., 2012).In practice, the atomic models of the resting state have either been refined by imposing inter-residue distances that are consistent with the experimentally derived constraints (Yarov-Yarovoy et al., 2006, 2012; Delemotte et al., 2010, 2011), or by explicitly modeling the side chains involved in the various cross-links or metal bridges themselves (Vargas et al., 2011; Henrion et al., 2012). Although these various models display high similarity, they all relied to varying degrees on computational “shortcuts” to obtain meaningful results about the VSD conformations within a reasonable computational time. Ultimately, the dream would be to “visualize,” atom-by-atom, how the channel moves as a function of time in response to a realistic membrane potential.This has now become possible in part by relying on the virtual reality provided by computer simulations. Computer trajectories “simulating” the effect of membrane hyperpolarization on a voltage-gated ion channel were generated by several research groups, with the goal of triggering deactivation to directly observe the conformational response and reorganization of the VSD (Treptow et al., 2004, 2009; Nishizawa and Nishizawa, 2008, 2009; Bjelkmar et al., 2009; Denning et al., 2009; Delemotte et al., 2011; Freites et al., 2012). More recently, Jensen et al. (2012) used long (hundreds of microseconds) MD simulations to visualize a complete spontaneous conformational transition of a voltage-gated K⁺ channel upon changes of the membrane potential. As in the previous simulations, large negative hyperpolarizing membrane potentials were applied (−750 mV) to shorten the time for the voltage-dependent transition toward the resting state within accessible computing time (although some simulations were also generated at −375 mV). The long MD simulations performed by Jensen et al. (2012) led to a computationally derived model of the resting-state conformation of the channel very similar to those deduced previously with different methods (Yarov-Yarovoy et al., 2006, 2012; Pathak et al., 2007; Bjelkmar et al., 2009; Delemotte et al., 2010, 2011; Khalili-Araghi et al., 2010; Schwaiger et al., 2011; Henrion et al., 2012) (Fig. 2). The results confirm and substantially strengthen the consensus from previous computational studies.Open in a separate windowFigure 2.Main elements of secondary structure of the VSD in the active and resting state. (A) The VSD in the active conformation (taken from the x-ray structure; PDB accession no. 3LUT). (B) A superposition of different models of the resting-state configuration of the VSD obtained by independent research teams (Delemotte et al., 2010; Schwaiger et al., 2011; Vargas et al., 2011; Jensen et al., 2012) using different constraints and methodologies. The four helices, S1 (gray), S2 (yellow), S3 (red), and S4 (blue), are displayed. The spheres correspond to the Cα atoms of E1 (Glu226) and E2 (Glu236) along S2, and R1 (Arg294) along S4.By several quantitative measures, all the mentioned atomic models of the resting state display a high degree of similarity, indicating that a consensus on the structure of the resting state and the mechanism of VSD function has emerged from the independent computational studies. The backbone Cα carbons of all the models lie within 3–4 Å root mean square deviation (RMSD) of one another (Table S1). This is comparable to the RMSD among the four VSDs of tetrameric structures obtained in various models. The RMSD among the four VSDs of the tetrameric structure of Delemotte et al. (2010, 2011) vary between 2.7 and 3.9 Å, and the RMSD between the four VSDs of the tetrameric structure of Jensen et al. (2012) vary between 1.3 and 3.4 Å. For comparison, the VSDs of the x-ray structures of the Kv1.2 and Kv1.2/Kv2.1 chimera display RMSDs of 1.5 Å for the same region. The largest deviations of any of the resting-state models from a hypothetical average configuration are <3 Å; the two main outliers are one subunit from Delemotte et al. (2010, 2011) and one subunit from Jensen et al. (2012) (Fig. S1).In all of the resting-state models, the S4 helix is predominantly rotated and translated inward along its main axis relative to the x-ray structures of the activated conformation, whereas the S1 and S2 helices retain their configuration. Averaging over all models, the absolute vertical displacement of S4 at the level of the Cα of the R1 position is ∼10 Å, with a spread of 3–4 Å (Table S1). There is some uncertainty in estimating the vertical translation of S4 because of the large structural fluctuations exhibited by the flexible VSDs. For example, the vertical position of R1 in the two x-ray structures of the active state of the K_V1.2 VSD differs by 2.6 Å (Long et al., 2005, 2007), and the net vertical displacement in the VSDs of the four subunits from Jensen et al. (2012) is 13.2, 16.2, 14.9, and 12.0 Å. All the resting-state models place the Cα of the R1 between the two acidic side chains E1 and E2 along the S2 helix, and R1 is also located above the highly conserved Phe located in the middle of S2. Most importantly, all of the resting-state models show the positive gating charge residues along S4 in position to either form salt bridges with acidic residues in the S1–S3 helices, or interact with the aqueous regions or the polar head groups. Similar conclusions have been obtained from the experiments on Na_V channels that combined structural modeling with disulfide cross-linking experiments (DeCaen et al., 2008, 2009, 2011; Yarov-Yarovoy et al., 2012).The term “resting state” in the above discussion is used to loosely describe the conformation in which S4 inhabits its most inward position. Upon careful consideration, however, “the” resting state is probably an oversimplified concept because it is likely that S4 does not withdraw to the same position in all channels as result of sequence variations, regardless of the applied membrane potential. Furthermore, several of these resting states of the VSD may favor the nonconducting closed state of the pore. Consistent with this idea, hyperpolarization of the membrane potential slows the activation of voltage-gated channels, a behavior known as the “Cole–Moore effect” (Cole and Moore, 1960), because the VSDs must move through more resting states before activation when they start from a more negative membrane potential. To highlight the structural differences among the various proposed resting-state models shown in Fig. 2 (right), it is useful to realign the different models with respect to the S1–S3 helices, which are the most stable structural elements. The set of models, displayed in Fig. S2, places the S4 helix at various depths relative to S1–S3, with a slight spread in the tilt angle of its main axis. The initial resting-state model of K_V1.2 (Yarov-Yarovoy et al., 2006) captured a resting state in which the S4 segment is not drawn as far inward as the models of Fig. 2 and which, therefore, likely represents an initial step toward activation. Models of a full range of resting states of the VSD extending to quite negative voltages have been developed for both K_V and Na_V channels (DeCaen et al., 2009, 2011; Delemotte et al., 2012; Henrion et al., 2012; Yarov-Yarovoy et al., 2012).The existence of multiple resting states is supported by a wide range of experiments, including analysis of multiple engineered metal bridges tracking successive states of the VSD (Henrion et al., 2012), noncovalent interactions between R2 and a tryptophan residue inserted in S1 (Lacroix et al., 2012), and disulfide cross-linking results of S4 gating charges with ion pair partners in S2 and S3 (DeCaen et al., 2009, 2011; Yarov-Yarovoy et al., 2012). It is expected that the multiple resting states of the VSDs of K_V and Na_V channels all stabilize the pore in its nonconducting closed conformation over a specific range of negative membrane potentials. Interestingly, a series of such resting states is observed in the long MD simulations of Jensen et al. (2012) as pauses in the inward movement of the VSD after the pore has closed. Thus, the conformations obtained from extremely long MD simulations are consistent with the resting state predicted by several independent studies using various computational approaches including knowledge-based structure prediction algorithms (Yarov-Yarovoy et al., 2006, 2012; Pathak et al., 2007) and MD simulations with experimental constraints (Delemotte et al., 2010; Vargas et al., 2011; Henrion et al., 2012). The final resting state from long MD simulations of Jensen et al. (2012) appears closest to the state reported in Delemotte et al. (2012), with the most inward position of S4. It is very satisfying that the conformational states are visited as the result of spontaneous transitions during long unbiased MD trajectories of the protein submitted to a negative membrane potential. The ability to simulate the spontaneous conformational transitions strengthens confidence in our current understanding of the physical forces and molecular interactions governing the voltage-gating process at the atomic level.

Mechanism of voltage-dependent activation

A consistent mechanistic perspective of voltage gating has emerged from these computational studies. The scenario that most accurately conveys the conformational change occurring within the VSD during activation as observed in these computations is the classical helical screw–sliding helix mechanism in which the S4 segment retains its helical conformation as it moves principally along its long axis (Catterall, 1986a,b; Guy and Seetharamulu, 1986). The gating charges are not directly exposed to the lipid hydrocarbon, and the S3–S4 helix-turn-helix does not move as a highly concerted structural motif across the membrane during voltage gating as proposed in the more recent paddle model (Jiang et al., 2003). Rather, sequential formation of salt bridges involving the gating residues plays an important role as proposed by Clay Armstrong (1981). Lastly, the concept of the “focused electric field” (Islas and Sigworth, 2001; Asamoah et al., 2003; Starace and Bezanilla, 2004), in which the spatial variation in the transmembrane potential affecting the gating charges of the VSD is concentrated over a narrow region that is considerably thinner than the full bilayer membrane, has been clarified by explicit calculations of the gating charge contributions based on all-atom MD simulations following two different approaches (Khalili-Araghi et al., 2010; Delemotte et al., 2011) and further supported by structural modeling studies (Yarov-Yarovoy et al., 2012). The gating charge calculations of Jensen et al. (2012) following the methodology of Khalili-Araghi et al. (2010) provided an additional confirmation of the concept of a focused electric field. Nonetheless, the S4 segment moves outward through the focused field, and the mechanism of voltage gating is not primarily a rearrangement in the transmembrane field as proposed in the transporter model (Chanda et al., 2005).The MD simulations of Jensen et al. (2012) showed that the ion-conducting pore closes before any of the four VSDs have undergone a transition to the most stable resting-state conformation. This sequence of events is consistent with kinetic models with discrete states developed long ago to describe voltage gating in the Shaker K⁺ channel (Bezanilla et al., 1994; Zagotta et al., 1994; Schoppa and Sigworth, 1998). According to these kinetic models, the first step involved in closing an activated channel is the closing of the pore domain, followed by the independent transitions of the four VSDs toward the resting state. Recent x-ray structures of bacterial Na_V channels provide examples of this intermediate state, showing a closed pore domain associated with VSDs in their activated conformation (Payandeh et al., 2011, 2012; Zhang et al., 2012).In summary, the major advances are that the resting-state conformation of the VSD reached by the long MD simulations is consistent with the results of numerous previous studies using different computational methods (Yarov-Yarovoy et al., 2006, 2012; Pathak et al., 2007; Bjelkmar et al., 2009; Delemotte et al., 2010, 2011; Khalili-Araghi et al., 2010; Schwaiger et al., 2011; Henrion et al., 2012), and that the sequence of events seen in the long simulations appears to be in qualitative accord with classical kinetic models of the voltage-gating process (Bezanilla et al., 1994; Zagotta et al., 1994; Schoppa and Sigworth, 1998). However, there is no experimental data at the large negative voltages used in the long simulations, and one must be cautious in trying to extrapolate the experimental time constants determined around −100 mV for ionic currents (Rodríguez and Bezanilla, 1996) and gating currents (Rodríguez et al., 1998).

Novel mechanistic hypotheses from MD simulations

Some novel ideas about the mechanism of voltage-dependent gating are suggested by the computational studies but do not yet have direct structural or experimental support. For example, sections of the S4 segment are observed in 3₁₀ helical conformation in x-ray crystal structures (Long et al., 2007; Clayton et al., 2008; Vieira-Pires and Morais-Cabral, 2010; Payandeh et al., 2011, 2012; Zhang et al., 2012). An intriguing suggestion from several simulation studies is the concept of a sequential dynamical transition to a 3₁₀ helical conformation for all or part of the S4 segment as it moves through the most hydrophobic region of the VSD (DeCaen et al., 2009, 2011; Khalili-Araghi et al., 2010; Schwaiger et al., 2011; Yarov-Yarovoy et al., 2012). Although the presence of some amount of 3₁₀ helical conformation is supported by available data (Villalba-Galea et al., 2008), the concept of a dynamic 3₁₀ transition of S4 during the voltage-gating process will require further experimental validation.Another suggestion from the long MD simulations is that the closure of the pore domain is driven by a rapid de-wetting transition taking place in the intracellular vestibule. A similar de-wetting process was previously found in voltage-driven simulations of an isolated pore domain, in the absence of the VSDs (Jensen et al., 2010). The de-wetting process results in a closed pore with a nearly dry central cavity. However, evidence that ions may be captured in the cavity of a closing Shaker K⁺ channel argues against a complete de-wetting (Baukrowitz and Yellen, 1996a,b; Ray and Deutsch, 2006). We note here also that the structures of the preopen and inactivated states of prokaryotic Na_V channels have closed, water-filled pores (Payandeh et al., 2011, 2012; Zhang et al., 2012). More experimental work is required to determine whether de-wetting drives pore closure or arises only at very negative membrane potentials.Lastly, the relationship of the voltage-gating transition displayed by the long MD simulations to the three major conformations of a VSD (resting, activated, and relaxed) observed in most of the S4-based VSDs (Villalba-Galea et al., 2008; Lacroix et al., 2011) remains uncertain. Moreover, the transitions observed in long MD simulations do not appear to reproduce all of the early components of the gating current (time constant of ∼10 µs) observed in Shaker K⁺ channels (Sigg et al., 2003). New experiments in which long MD simulations and gating current measurements are made in parallel on the same channel will give more insight into these issues.

Confidence in the computational results

Several models derived from a combination of experimental data and computations, produced from different approaches, have converged to yield a low resolution picture of the resting-state conformation, defined within ∼3–4 Å RMSD (Fig. 2, right). Upon a closer look, fine differences can be noted among the various models (Fig. S2), which points to the concept of multiple resting states in which the segment S4 is drawn to different depths toward the intracellular side. The broad agreement among the various computational methods, most likely, is not fortuitous, and the picture emerging represents a genuine advance in understanding voltage-gated channels. The implication, if the computational results are to be trusted, is that many of the apparently conflicting measures about voltage gating can be resolved. Nevertheless, some might argue that the controversy about the resting-state conformation of the VSD will remain until an experimental x-ray structure becomes available. In this context, it is important to note that all structures are models, even x-ray crystal structures. However, structural models from x-ray crystallography rely on a huge amount of experimental data and are derived from rigorously established procedures that have been extensively tested and cross-validated. It is expected that experimental structure determination will increasingly rely on sophisticated computational modeling to complement low resolution data (Chen et al., 2007; Trabuco et al., 2008; Brunger et al., 2012). In the early 1980s, NMR structures were considered tentative models until it was demonstrated that the results were consistent with x-ray crystallography (Billeter et al., 1989). What ultimately matters is the quantifiable level of confidence that can be attributed to a structural model. As the methodologies become more and more reliable and consistent, computational modeling will play an increasingly important role in structural biology (DiMaio et al., 2011; Lange et al., 2012). The present situation, in which the proposed models of the VSD (Fig. 2) are supported by such a wide range of computational approaches applied by different investigators, is unprecedented to our knowledge. For this reason, our view is that one may be (cautiously) optimistic that the resting-state structures of Fig. 2 and the related structures in other computational papers cited here are close (within ∼3–4 Å RMSD) to reality. Notably, this accuracy can be predicted from the distribution of models themselves (even the early ones).At this point it is prudent to sound a note of caution. The progress documented in this Viewpoint on understanding structural aspects of a membrane-bound channel protein has been made possible by using novel computational methodologies and an empirical potential energy (force field) that subsumes polarization and nuclear quantum effects in an average fashion. Although MD simulation articles often imply that every in silico detail from the trajectory is real, it is useful to remind ourselves that this is not necessarily the case. Experience indicates that current approximations are more successful in predicting conformational states than transition rates. The reason is that the overall topology of the potential energy surface, with its wells and barriers, is more or less correct, even though the relative depths of the wells and heights of the barriers may be imperfect. As a consequence, the order in which the chain of events takes place in a simulation during a complex conformational transition may not reflect reality, as some parts may undergo transitions that are too slow, whereas other parts undergo transitions that are too fast. Therefore, although a strong consensus is emerging on the nature of the conformation of the resting state, the dynamic properties of the gating process require more scrutiny. This will be challenging notwithstanding the massively increased length of recent MD trajectories.

Conclusion

A clear consensus on the mechanism of voltage-dependent gating is emerging from various studies based on a wide range of computational and experimental methods. This consensus, which is to be celebrated, highlights the increasingly important role of computational modeling in linking molecular structure to biological function by supplementing missing information. It is important, however, to remain prudent in assessing the significance of details and features of the computationally derived models that have not yet been experimentally validated. Even if the resting-state conformation of the VSD reached by the simulations is correct, and the sequence of events is in accord with classical kinetic models of the voltage-gating process, it is possible that the rates of the individual processes is differentially affected by insufficient sampling, force field inaccuracies, and the large membrane potential typically applied so far. Nevertheless, it is encouraging to note that, despite their inherently approximate nature, current computational models can provide meaningful answers to important questions about complex biomolecular systems. Further studies using computational methods in concert with structure–function experiments seem likely to soon reveal the missing details of VSD function.

Online supplemental material

Table S1 provides all the data about minimum global RMSD of the VSDs using best pairwise alignments for all models. Fig. S1 shows the deviation of backbone atom of each model relative to the average. Fig. S2 shows a superposition of all the VSD models in the resting-state configuration aligned with respect to S1–S3 helices. Fig. S3 presents a quantitative structural comparison of all the VSD models with the crystal structures. Video 1 includes an animation showing all the available VSD models rotating in superposition. The online supplemental material is available at http://www.jgp.org/cgi/content/full/jgp.201210873/DC1.     

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.     

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.]     

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.     

Identification of a Novel Abscisic Acid-Regulated Farnesol Dehydrogenase from Arabidopsis

In Arabidopsis (Arabidopsis thaliana), farnesylcysteine is oxidized to farnesal and cysteine by a membrane-associated thioether oxidase called farnesylcysteine lyase. Farnesol and farnesyl phosphate kinases have also been reported in plant membranes. Together, these observations suggest the existence of enzymes that catalyze the interconversion of farnesal and farnesol. In this report, Arabidopsis membranes are shown to possess farnesol dehydrogenase activity. In addition, a gene on chromosome 4 of the Arabidopsis genome (At4g33360), called FLDH, is shown to encode an NAD⁺-dependent dehydrogenase that oxidizes farnesol more efficiently than other prenyl alcohol substrates. FLDH expression is repressed by abscisic acid (ABA) but is increased in mutants with T-DNA insertions in the FLDH 5′ flanking region. These T-DNA insertion mutants, called fldh-1 and fldh-2, are associated with an ABA-insensitive phenotype, suggesting that FLDH is a negative regulator of ABA signaling.Isoprenylated proteins are modified at the C terminus via cysteinyl thioether linkage to either a 15-carbon farnesyl or a 20-carbon geranylgeranyl group (Clarke, 1992; Zhang and Casey, 1996; Rodríguez-Concepción et al., 1999; Crowell, 2000; Crowell and Huizinga, 2009). These modifications mediate protein-membrane and protein-protein interactions and are necessary for the proper localization and function of hundreds of proteins in eukaryotic cells. In Arabidopsis (Arabidopsis thaliana), the PLURIPETALA (PLP; At3g59380) and ENHANCED RESPONSE TO ABA1 (At5g40280) genes encode the α- and β-subunits of protein farnesyltransferase (PFT), respectively (Cutler et al., 1996; Pei et al., 1998; Running et al., 2004). These subunits form a heterodimeric zinc metalloenzyme that catalyzes the efficient transfer of a farnesyl group from farnesyl diphosphate to protein substrates with a C-terminal CaaX motif, where “C” is Cys, “a” is an aliphatic amino acid, and “X” is usually Met, Gln, Cys, Ala, or Ser (Fig. 1). The PLP and GERANYLGERANYL-TRANSFERASE BETA (At2g39550) genes encode the α- and β-subunits of protein geranylgeranyltransferase type 1 (PGGT1), respectively (Running et al., 2004; Johnson et al., 2005). These subunits form a distinct heterodimeric zinc metalloenzyme that catalyzes the efficient transfer of a geranylgeranyl group from geranylgeranyl diphosphate to protein substrates with a C-terminal CaaL motif, where “C” is Cys, “a” is an aliphatic amino acid, and “L” is Leu. A third protein prenyltransferase, called protein geranylgeranyltransferase type II or RAB geranylgeranyltransferase, catalyzes the dual geranylgeranylation of RAB proteins with a C-terminal XCCXX, XXCXC, XXCCX, XXXCC, XCXXX, or CCXXX motif, where “C” is Cys and “X” is any amino acid. However, RAB proteins must be associated with the RAB ESCORT PROTEIN to be substrates of RAB geranylgeranyltransferase. Plant protein prenylation has received considerable attention in recent years because of the meristem defects of Arabidopsis PFT mutants and the abscisic acid (ABA) hypersensitivity of Arabidopsis PFT and PGGT1 mutants (Cutler et al., 1996; Pei et al., 1998; Running et al., 1998, 2004; Johnson et al., 2005).Open in a separate windowFigure 1.Proposed metabolism of farnesal and farnesol as it relates to protein prenylation. The portion of the cycle shown in red is the subject of this article.Proteins that are prenylated by either PFT or PGGT1 undergo further processing in the endoplasmic reticulum (Crowell, 2000; Crowell and Huizinga, 2009). First, the aaX portion of the CaaX motif is removed by proteolysis (Fig. 1). This reaction is catalyzed by one of two CaaX endoproteases, which are encoded by the AtSTE24 (At4g01320) and AtFACE-2 (At2g36305) genes (Bracha et al., 2002; Cadiñanos et al., 2003). Second, the prenylated Cys residue at the new C terminus is methylated by one of two isoprenylcysteine methyltransferases (Fig. 1), which are encoded by the AtSTE14A (At5g23320) and AtSTE14B (ICMT; At5g08335) genes (Crowell et al., 1998; Crowell and Kennedy, 2001; Narasimha Chary et al., 2002; Bracha-Drori et al., 2008). A specific isoprenylcysteine methylesterase encoded by the Arabidopsis ICME (At5g15860) gene has also been described, demonstrating the reversibility of isoprenylcysteine methylation (Deem et al., 2006; Huizinga et al., 2008).Like all proteins, prenylated proteins have a finite half-life. However, unlike other proteins, prenylated proteins release farnesylcysteine (FC) or geranylgeranylcysteine (GGC) upon degradation. Mammals possess a prenylcysteine lyase enzyme that catalyzes the oxidative cleavage of FC and GGC (Zhang et al., 1997; Tschantz et al., 1999; Tschantz et al., 2001; Beigneux et al., 2002; Digits et al., 2002). This FAD-dependent thioether oxidase consumes molecular oxygen and generates hydrogen peroxide, Cys, and a prenyl aldehyde product (i.e. farnesal or geranylgeranial). In Arabidopsis, a similar lyase exists. However, the Arabidopsis enzyme, which is encoded by the FCLY (At5g63910) gene, is specific for FC (Fig. 1; Crowell et al., 2007; Huizinga et al., 2010). GGC is metabolized by a different mechanism.Plant membranes have been shown to contain farnesol kinase, geranylgeraniol kinase, farnesyl phosphate kinase, and geranylgeranyl phosphate kinase activities (Fig. 1; Thai et al., 1999). These membrane-associated kinases differ with respect to nucleotide specificity, suggesting that they are distinct enzymes (i.e. farnesol kinase and geranylgeraniol kinase can use CTP, UTP, or GTP as a phosphoryl donor, whereas farnesyl phosphate kinase and geranylgeranyl phosphate kinase exhibit specificity for CTP as a phosphoryl donor). However, it remains unclear if farnesol kinase is distinct from geranylgeraniol kinase or if farnesyl phosphate kinase is distinct from geranylgeranyl phosphate kinase. Nonetheless, it is clear that these kinases convert farnesol and geranylgeraniol to their monophosphate and diphosphate forms for use in isoprenoid biosynthesis, including sterol biosynthesis and protein prenylation.Because plants have the metabolic capability to generate farnesal from FC and farnesyl diphosphate from farnesol, we considered the possibility that plant membranes also contain an oxidoreductase capable of catalyzing the reduction of farnesal to farnesol and/or the oxidation of farnesol to farnesal (Fig. 1; Thai et al., 1999; Crowell et al., 2007). To date, the only reports of such an oxidoreductase are from the corpora allata glands of insects, where it participates in juvenile hormone synthesis, and black rot fungus-infected sweet potato (Ipomoea batatas; Baker et al., 1983; Inoue et al., 1984; Sperry and Sen, 2001; Mayoral et al., 2009). Insect farnesol dehydrogenase is an NADP⁺-dependent oxidoreductase that is encoded by a subfamily of short-chain dehydrogenase/reductase (SDR) genes (Mayoral et al., 2009). Farnesol dehydrogenase from sweet potato is a 90-kD, NADP⁺-dependent homodimer with broad specificity for prenyl alcohol substrates and is induced by wounding and fungus infection of potato roots (Inoue et al., 1984).Here, we extended previous work in which [1-³H]FC was shown to be oxidized to [1-³H]farnesal, and [1-³H]farnesal reduced to [1-³H]farnesol, in the presence of Arabidopsis membranes (Crowell et al., 2007). The reduction of [1-³H]farnesal to [1-³H]farnesol was abolished by pretreatment of Arabidopsis membranes with NADase, suggesting that sufficient NAD(P)H is present in Arabidopsis membranes to support the enzymatic reduction of farnesal to farnesol. In this report, we demonstrate the presence of farnesol dehydrogenase activity in Arabidopsis membranes using [1-³H]farnesol as a substrate. Moreover, we identify a gene on chromosome 4 of the Arabidopsis genome (At4g33360), called FLDH, that encodes an NAD⁺-dependent dehydrogenase with partial specificity for farnesol as a substrate. FLDH expression is repressed by exogenous ABA, and fldh mutants exhibit altered ABA signaling. Taken together, these observations suggest that ABA regulates farnesol metabolism in Arabidopsis, which in turn regulates ABA signaling.     

Structural basis of the super- and hyper-relaxed states of myosin II

Super-relaxation is a state of muscle thick filaments in which ATP turnover by myosin is much slower than that of myosin II in solution. This inhibited state, in equilibrium with a faster (relaxed) state, is ubiquitous and thought to be fundamental to muscle function, acting as a mechanism for switching off energy-consuming myosin motors when they are not being used. The structural basis of super-relaxation is usually taken to be a motif formed by myosin in which the two heads interact with each other and with the proximal tail forming an interacting-heads motif, which switches the heads off. However, recent studies show that even isolated myosin heads can exhibit this slow rate. Here, we review the role of head interactions in creating the super-relaxed state and show how increased numbers of interactions in thick filaments underlie the high levels of super-relaxation found in intact muscle. We suggest how a third, even more inhibited, state of myosin (a hyper-relaxed state) seen in certain species results from additional interactions involving the heads. We speculate on the relationship between animal lifestyle and level of super-relaxation in different species and on the mechanism of formation of the super-relaxed state. We also review how super-relaxed thick filaments are activated and how the super-relaxed state is modulated in healthy and diseased muscles.

The super-relaxed state of myosinAnimal life is characterized by the constant need for food, which provides the raw materials for making ATP used in the body’s energy-requiring processes. A substantial amount of energy is expended by skeletal muscle, which in humans amounts to 30–40% of body mass (Janssen et al., 2000). During contraction, ATP is rapidly consumed by the molecular motor, myosin II, as it pulls on actin filaments to produce force and movement. But ATP is also used at a significant basal rate even in the resting state. Producing ATP is costly for the cell, so there is a substantial evolutionary advantage to minimizing its waste. Animals have adapted to this requirement by evolving a mechanism that reduces the consumption of ATP to minimal levels in relaxed (RX) muscle. This mechanism is known as super-relaxation or the super-relaxed state (SRX).SRX is a biochemical state in which ATP turnover by a portion of the myosin heads in the RX state of muscle thick filaments is much slower (∼10 times) than that of myosin II molecules in solution (Stewart et al., 2010; Cooke, 2011; Nag and Trivedi, 2021). This inhibited state, in equilibrium with a faster (RX) state, was suggested by early studies showing that the metabolic rate of live, resting muscle was much less than that expected from the ATP turnover rate of purified myosin (Ferenczi et al., 1978) and similarly that ATPase activity of RX myofibrils was also lower than expected from isolated myosin (Myburgh et al., 1995). A slow rate of ATP turnover was later directly demonstrated in studies of skinned muscle fibers (Stewart et al., 2010). The single ATP turnover rate is typically measured as the rate of binding of fluorescent ATP (mantATP) to, or release from, myosin heads in solution or in skinned fiber bundles (Hooijman et al., 2011; McNamara et al., 2015); conceptually, because P_i release is rate limiting, the apparent rates of ATP binding and ADP release effectively reveal the overall ATP turnover rate. Experimental curves are generally interpreted in terms of two exponentials, one with a slow rate (SRX) and the other with a faster but still slow (RX) rate of ATP use (Cooke, 2011), although additional exponentials can sometimes be resolved (Hooijman et al., 2011; Naber et al., 2011).The SRX state is ubiquitous and thought to be fundamental to muscle function, acting as a mechanism for parking and switching off—like a car—energy-consuming myosin motors when they are not being used. It has been detected in all muscles where it has been studied: vertebrate (including human) and invertebrate, fast and slow skeletal, and cardiac muscle (Stewart et al., 2010; Cooke, 2011; Hooijman et al., 2011; Naber et al., 2011; McNamara et al., 2015; Phung et al., 2020). A highly inhibited rate of ATP turnover also characterizes the switched-off state of vertebrate smooth muscle and nonmuscle myosin II molecules (Cross et al., 1988). SRX plays a critical role in muscle energetics, conserving ATP in resting as well as contracting muscles and providing a reserve of myosin heads for enhanced contractility in cardiac muscles (Cooke, 2011; Hooijman et al., 2011; Brunello et al., 2020; Ma et al., 2021b).What are the key features of the SRX state?The SRX state has two main parameters: (1) the rate of ATP use (i.e., ATP turnover rate, often expressed as its inverse, the ATP turnover time) and (2) the fraction of molecules exhibiting this rate. Both parameters are essential to understanding the significance of SRX in the energy balance of muscle: myosin heads with a very slow rate would be of little consequence if not present in substantial numbers. The rate of ATP use has been attributed to the specific conformation of the myosin head, which can impede, or not, the release of the products of ATP hydrolysis (ADP and P_i; Anderson et al., 2018), and the fraction of heads with this rate in muscle appears to be related to the intra- and intermolecular interactions that the heads undergo in thick filaments, as discussed below.ATP turnover by myosin heads can vary by six orders of magnitude, depending on the muscle and its state of activity (Cooke, 2011; Naber et al., 2011). These fall into four main groups, which, on a logarithmic scale, differ internally by small amounts, but by an order of magnitude or more between groups (Fig. 1 A). The fastest use of ATP is in contracting muscle (rate ≈ 10 s⁻¹), where actin activates the release of hydrolysis products in a process coupled to the power stroke of the myosin head. The advantages of force and movement are paid for by a rapid consumption of ATP. RX heads use ATP at least 100 times more slowly: some at the RX rate, similar to that of purified myosin in solution (<0.1 s⁻¹), and others at an SRX rate, 10 times slower (Fig. 1, A and B). As might be predicted from the two parameters of the SRX, nature has evolved different ways of saving energy in this state: decreasing the ATP turnover rate, increasing the fraction with the low rate, or a combination of the two. These strategies appear adapted to the different systems in which they are found. The SRX rate typically occurs in 50% or more of the heads in a thick filament, the balance being RX heads (Cooke, 2011). In certain muscles, there is an even slower rate (10 times slower than SRX), which we refer to as “hyper-relaxed” (HRX), present together with SRX and RX heads (Naber et al., 2011).Open in a separate windowFigure 1.ATP turnover rates of myosin and proposed relation to head organization. (A) Logarithmic plot of rates, varying from very slow (hyper-relaxed), to slow (super-relaxed), to fast (disordered-relaxed), to very fast (actin-activated). (B) Expanded plot of relaxed rates and proposed relation to head configuration and interactions. The cartoons show the IHM as found in single molecules (upper) and thick filaments (lower); in filaments, only molecules in a single crown are shown. The colored ellipses are the interacting motor domains. The smaller, gray domains are the ELCs and RLCs, and the gray lines are the proximal region of the myosin tail (S2). Molecules with a sufficient length of S2 (25 heptads [25 hep]; ∼250 Å) form an IHM, while single heads and two-headed molecules with only two heptads of tail show no interactions. Molecules in filaments have SRX and DRX turnover rates similar to those of isolated molecules (25-heptad IHM; cf. vertebrate IHM) or more inhibited (HRX) rates. Heads turn over ATP at fast (F; noninteracting), slow (S; interacting), or very slow (VS; more interactions) rates. Slow (attached) FHs of the IHM can detach and sway out (red, double-headed arrows) and, while detached, can equilibrate between the slow and fast conformations, similar to S1 (blue, reversible arrows). The “tail” in tarantula and the uncolored IHM in scallop provide additional, intermolecular interactions leading to the very slow rate; this rate is also seen in 10S myosin through intramolecular interactions with segment 2 of its own tail. The different lengths of the bars in A correspond to the lengths in B, which are drawn to include the range of rates for HRX, SRX, and DRX; the key point is the close clustering of rates within these groups and the large gaps between them.What is the structural basis of the SRX state?The early finding that myosin filaments in muscle had a slower ATP turnover rate than myosin II molecules in solution led to the idea that interactions of heads possible in the polymer (e.g., with the thick filament backbone), but not in the monomer, inhibited their activity (Myburgh et al., 1995; Stewart et al., 2010; Cooke, 2011). The first study to unequivocally show head organization in a thick filament indeed revealed such interactions—of heads with the backbone, with each other within a molecule, and with each other between molecules (Woodhead et al., 2005). Of particular note was a folding back of the two heads onto the proximal part of the tail (subfragment 2 [S2]), with which they interacted, and intramolecular interaction between the heads through their motor domains. This structure became known as the “interacting-heads motif” (IHM; Fig. 1 B, dotted box; Fig. 2 B; Woodhead et al., 2005; Alamo et al., 2008). Such interaction between heads had first been seen in isolated myosin molecules (Wendt et al., 2001; Burgess et al., 2007) and was thought to inhibit their activity. The two heads were called “blocked” and “free” (BH and FH, respectively), referring to their actin-binding capability (Wendt et al., 2001). In this model, actin binding by the BH would be blocked by interaction of its actin-binding site with the FH. While the FH actin-binding site was exposed, movement of its converter domain, needed for ATP product release, would be inhibited by binding to the BH (Wendt et al., 2001). The overall result would be inhibition of activity of both heads, but by different mechanisms. Interaction of the folded-back heads with S2 would further inhibit the motions required for ATPase activity (Woodhead et al., 2005).Open in a separate windowFigure 2.Structural basis of SRX. (A) Proposed conformations of myosin heads (S1), in equilibrium with each other, underlying SRX (left, bent) and DRX (right, straight) ATP turnover rates (Anderson et al., 2018). MD, motor domain. (B) IHM of cardiac HMM showing BH and FH (based on Protein Data Bank accession no. 5TBY). Ellipses show regions of interaction between BH and FH motor domains (black ellipse), FH and S2 (black circle), and BH and S2 (white ellipse; interaction occurs on rear side of BH). (C) Thick filament (tarantula; EM Data Bank accession no. 1950) showing IHMs lying along four coaxial helices (three on front marked with arrows) creating intermolecular interactions (ellipses) between FH of one IHM and regulatory domain of IHM above. M-line would be at the top of the image. The reconstruction shows the average positions of the heads in the filament; however, the FHs are thought to be dynamic, leaving and returning to the IHM (Fig. 1 B; see text). Models in this figure were created with UCSF Chimera (Pettersen et al, 2004).The IHMs in RX thick filaments are organized in helical arrays, with intermolecular interactions of IHMs with each other along the helices as well as with the filament backbone (Woodhead et al., 2005; Zoghbi et al., 2008; Zhao et al., 2009; Pinto et al., 2012; Woodhead et al., 2013; Sulbarán et al., 2015). Helical ordering requires the closed conformation of the myosin heads (Xu et al., 2003; Zoghbi et al., 2004; Xu et al., 2009) and is thought to be a signature of the IHM. It can be disrupted in multiple ways (increased salt level, phosphorylation of the myosin regulatory light chains [RLCs], substitution of GTP or ADP for ATP, lowering of temperature), and this is accompanied by reduction of the SRX state in each case (Stewart et al., 2010; Cooke, 2011). Conversely, conditions that enhance the IHM, such as treatment with the myosin inhibitor blebbistatin (Zhao et al., 2008; Xu et al., 2009; Fusi et al., 2015), enhance the SRX (Wilson et al., 2014; Fusi et al., 2015). These correlations led to the view that IHMs in the ADP.Pi prepowerstroke state (Xu et al., 1999), organized in helices and bound to the core of the thick filament, may be the structural basis of the SRX state (Stewart et al., 2010; Cooke, 2011; Wilson et al., 2014; Fusi et al., 2015; Alamo et al., 2016).Despite this suggestive evidence, however, recent single ATP turnover measurements of myosin constructs have shown that slow turnover of ATP occurs not only in thick filaments but also, to a small extent (∼10–20% of molecules), in isolated myosin heads (S1; Anderson et al., 2018; Rohde et al., 2018; Gollapudi et al., 2021b; the earlier, steady-state solution ATPase observations were not capable of revealing these low rates). Thus, neither the filamentous form of myosin nor the IHM is required for the SRX rate of ATP turnover, as originally thought. It has been suggested, instead, that myosin heads in solution exist in an equilibrium between a strongly bent (“closed”) conformation (in which the lever arm is tilted more toward the prestroke direction similar to that in the IHM structure; 10–20% of molecules) and a more extended (“open”) structure (Anderson et al., 2018) and that inhibition of phosphate release and thus ATP turnover by the closed structure could account for the observation of a small level of SRX in S1 (Figs. 1 B and 2 A). If this is the case, what is the role of the IHM in SRX?Different degrees of SRX correlate with different levels of head organizationMyosin heads can exist in several structural forms: single heads, two-headed myosin molecules or constructs, and the polymeric myosin filaments found in muscle. Studies show that SRX increases (greater inhibition or greater number of inhibited molecules) as myosin heads are incorporated into more complex structures. As mentioned, ∼10–20% of isolated myosin heads in solution have a slow rate (SRX) and are thought to be in equilibrium with the balance of heads having a turnover rate similar to the conventionally measured ATPase (RX heads; Anderson et al., 2018; Rohde et al., 2018; Gollapudi et al., 2021b). When myosin heads are present in two-headed constructs containing 25 heptads of tail (∼250 Å; Figs. 1 B and 2 B) or full-length S2, similar SRX and RX rates are observed, but the fraction of SRX heads increases to ∼25–30% (at physiological ionic strength; Anderson et al., 2018; Rohde et al., 2018; Gollapudi et al., 2021b), suggesting that the presence of two heads and/or the tail stabilizes the SRX head structure. Importantly, if the tail is short (only two heptads; Fig. 1 B), the rates and fractions are similar to those of S1 (Anderson et al., 2018), demonstrating the importance of the tail in creating the SRX. Modeling and experiments show that 25 heptads (or more) of tail is long enough for molecules to form an IHM with folded-back, interacting FHs and BHs (Figs. 1 B and 2 B; Anderson et al., 2018). Head–head and head–tail interactions would stabilize the heads in their inhibited conformations (Fig. 2 B), consistent with the increase in SRX fraction. The two-heptad-long tail is too short for these interactions (Fig. 1 B), and the molecule does not show evidence of head interactions (Nag et al., 2017): the heads behave like isolated S1 in solution, with only ∼10% having the SRX rate (Fig. 1 B; Anderson et al., 2018). We conclude that the stabilizing interactions in the IHM involving the tail and the two heads increase the number of heads in the SRX conformation.Studies of whole muscle fibers (vertebrate fast skeletal) show that when myosin molecules are in native thick filaments in their helically ordered IHM conformations (Zoghbi et al., 2008; AL-Khayat et al., 2013), the RX and SRX rates are again similar to those of S1 (Fig. 1 B), but the fraction of SRX now reaches 60–75% (Stewart et al., 2010; Cooke, 2011). A similar, greatly increased fraction of SRX heads is found in slow skeletal and cardiac muscle fibers and myofibrils (Stewart et al., 2010; Hooijman et al., 2011; McNamara et al., 2017; Nelson et al., 2020; Gollapudi et al., 2021a). In filaments, IHMs undergo intermolecular interactions that cannot occur with single molecules (Zoghbi et al., 2008; AL-Khayat et al., 2013). These include interactions between heads of different IHMs along the myosin helices (Fig. 2 C), interaction of heads with tails in the filament backbone, and interaction with myosin-binding protein C (MyBP-C) and titin, lying on the filament surface (Nag et al., 2017). We suggest that these interactions further stabilize the FHs and BHs of the IHMs, without significantly affecting their conformation, leading to the high percentage of SRX in filaments (Anderson et al., 2018).These observations are consistent with the concept suggested earlier that the SRX rate is a consequence of a specific conformation of myosin heads, which inhibits ATP turnover, and with stabilization of this conformation by intra- and intermolecular interactions of the heads (Anderson et al., 2018). When heads are attached to each other in myosin constructs with a sufficient length of tail, forming the IHM, the inhibited conformation is stabilized by intramolecular interactions of the two heads with each other and with the myosin tail, approximately doubling (25–30%) the SRX fraction found in single heads (10–20%). When IHMs are incorporated into thick filaments, the inhibited heads are further stabilized by intermolecular interactions, with a further doubling of the SRX fraction to 60–75%. Thus, while the IHM is not a requirement for the SRX state, in real muscle, it strongly enhances it.Are the RX and SRX rates associated with specific heads in the IHM? It is known experimentally that the BH is more stably associated with the IHM than the FH. EM images of smooth muscle heavy meromyosin (HMM; a soluble, proteolytic fragment of myosin containing the two heads and the proximal third of the tail) show molecules where the BH is attached to S2 but the FH has dissociated (Burgess et al., 2007). This and studies of tarantula thick filaments (Brito et al., 2011; Sulbarán et al., 2013) suggest that the FH can dissociate from the IHM and become mobile (“swaying”; Fig. 1 B, red double-headed arrows), with a duty ratio that defines the fraction of time spent in the dissociated state (Alamo et al., 2017). When unconstrained in this way, the FH presumably acts essentially like S1 in solution, equilibrating between its minor SRX (bent) and its predominant RX (straight) rate of ATP turnover (Fig. 1 B, reversible blue arrows). Thus, the more stable BH would account for most of the SRX rate and the dissociated FH for the faster, RX rate. When the FH is docked in the IHM, stabilized in its inhibited form by interactions with the BH and S2 in the IHM, it may slow to approximately the BH SRX rate (Fig. 1 B; Alamo et al., 2016). The BH and docked FH may have similar enough rates that they are not resolved by the two-exponential analysis; there is in fact some evidence that additional exponentials give an improved fit (Hooijman et al., 2011), but this has not been explored in detail. The idea that some heads can move freely for a portion of the time, leading to their disordering in thick filaments, and the correlation of disorder with the RX rate, has led to the concept of the “disordered relaxed state” (DRX; Wilson et al., 2014). Thus, heads in thick filaments are typically referred to as SRX or DRX.What is the structural basis of the HRX state?Studies of invertebrate (tarantula) striated muscle show that the SRX state in some species can be enhanced by a further (10 times) slowing of the ATP turnover rate compared with the SRX (Fig. 1) and that this occurs in up to 50% of the heads (Naber et al., 2011). We refer to this as the HRX state. The HRX state has not been observed in vertebrate striated muscle thick filaments or myosin molecules. Is there a structural explanation for the greater inhibition of ATP turnover in tarantula muscle? The cryo-EM reconstruction of tarantula filaments shows that the IHMs in this species are arranged in four coaxial helices, with crowns of four IHMs spaced 145 Å apart axially (Woodhead et al., 2005). Each IHM shows the conventional structure of two heads interacting with each other through their motor domains and with the proximal portion of the myosin tail. Importantly, the reconstruction also suggests an additional key interaction that is apparently absent in vertebrate thick filaments (see below). S2, after leaving its IHM, travels along the filament toward the bare zone, passing near the next IHM along the filament axis, through a groove formed by the BH SRC homology 3 (SH3) and converter domains (Fig. 3 A; and Fig. 4, A and B; Woodhead et al., 2005). This location suggests that S2 could sterically interfere with movement of the BH converter region of this IHM that is required for product release. It could thus act as an additional constraint on the already inhibited conformation of the BH that further inhibits ATP turnover (each BH would thus interact with two S2s, its own and that from the axially adjacent IHM). In this case, we suggest that intermolecular interaction does not simply stabilize the inhibited head conformation but creates a new and additional physical barrier to motions of the BH required for ATPase activity, leading to the highly inhibited HRX turnover rate. The reconstruction shows that only the BHs are affected in this way, which could directly explain the finding that 50% of heads are in the HRX state (Naber et al., 2011).Open in a separate windowFigure 3.Comparison of intermolecular interactions in 3-D reconstructions of different thick filaments. Reconstructions are fitted with IHM atomic models (Protein Data Bank accession nos. 3JBH or 5TBY) in each case. All thick filaments (except insect flight muscle; Hu et al., 2016) show interactions between IHMs along the helical tracks (arrows), involving the FH motor domain at one level and the BH lever arm at the next. These interactions are the same at every level in tarantula and scallop (true helical structures) but different at different levels in vertebrates, which are quasi-helical. Additional interactions vary between species. (A) Tarantula shows interaction of S2 from one crown of heads with SH3 and converter domains of the next level (circles; see Fig. 4, A and B). IHM (Protein Data Bank accession no. 3JBH) was fitted to four levels of heads in cryo-EM reconstruction (EMD accession no. 1950; gray). (B) Similar fitting of IHM (Protein Data Bank accession no. 5TBY) to vertebrate (human) cardiac negative stain reconstruction of C-zone (EMD accession no. 2240) shows well-ordered IHMs at two of every three levels of heads (strong map density for pink and cyan IHMs), while the third level (yellow) is poorly ordered (weak map density, suggesting substantial IHM mobility; Zoghbi et al., 2008; AL-Khayat et al., 2013). S2 cannot be fitted unambiguously due to low resolution of map and lack of internal detail with negative stain; within these limitations, there is no obvious S2–head interaction between crowns (circles; see Video 1; cf. Video 2 for 3-D view). (C) In scallop cryo-EM reconstruction, S2 is not resolved, but the tight azimuthal crowding of IHMs around the circumference at each crown suggests potential intermolecular interaction between the SH3 domain of a BH and the motor domain of the neighboring FH (circles). Filaments are oriented with M-line at top; their different symmetries (fourfold, threefold, and sevenfold rotational symmetry, respectively) cause the varying views of the IHMs in the different filaments. All reconstructions are at the same scale. The human has a smaller diameter due to radial shrinkage occurring during negative staining and to the smaller number of molecules (n = 3) at each level. The scallop and tarantula cryo-reconstructions also have different diameters: scallop has seven molecules at each level forming a shell above the filament backbone (Woodhead et al., 2013), while tarantula has four molecules at each level, closer to the backbone (Woodhead et al., 2005), leading to a smaller diameter. Models in this figure were created with UCSF Chimera (Pettersen et al, 2004).Open in a separate windowFigure 4.Comparison of tail interactions with the SH3 and converter domains in atomic models of tarantula thick filaments and 10S smooth muscle myosin. (A and B) Tarantula thick filament (Protein Data Bank accession no. 3JBH). (C and D) 10S myosin (Protein Data Bank accession no. 6XE9). (A and C) Front views at same scale. (B and D) End views (same scale) obtained by rotating A and C 90° around the x axis so that pink IHM in A is closest to viewer (i.e., looking toward M-line). S2 in tarantula filament and segment 2 (Seg2) in 10S myosin both pass through a groove formed by the SH3 and converter (Cnv) domains of the BH motor domain (MD). The looser fit to the groove in tarantula may be due to the lower resolution of the reconstruction used to obtain the atomic model (20 Å resolution; cf. 4.3 Å for 10S myosin). Models in this figure were created with UCSF Chimera (Pettersen et al, 2004).Several observations support this structural interpretation of the HRX state. Vertebrate thick filaments have a symmetry and organization of myosin heads that is different from those in tarantula. Although no cryo-EM reconstruction of vertebrate filaments is available, two negative stain reconstructions of the C-zone (the middle section of each filament half, containing MyBP-C) clearly show the well-known quasi-helical arrangement of myosin heads characteristic of vertebrates (Zoghbi et al., 2008; AL-Khayat et al., 2013). Fitting of the IHM into the density map reveals well-ordered IHMs at two of every three levels of heads, while the third level is relatively disordered (Zoghbi et al., 2008; AL-Khayat et al., 2013). S2 cannot be fitted with certainty due to the low resolution of the map and ambiguities of negative stain; however, there is no obvious interaction of S2 from one crown with heads in the next (Fig. 3 B; Video 1; cf. Video 2). Correspondingly, vertebrate filaments do not exhibit an HRX state. There is also no HRX state in isolated S1- or S2-headed myosin constructs forming the IHM (Anderson et al., 2018; Rohde et al., 2018). These observations are consistent with the conclusion that in tarantula, intermolecular interaction with S2 from the neighboring crown is responsible for the HRX state. Note that the “ultra-relaxed” state induced in myosin by the inhibitor blebbistatin (Gollapudi et al., 2021a) has a very different origin from the HRX—the former due to internal stabilization of switch 2 in the myosin head in the closed state, inhibiting phosphate release (Zhao et al., 2008), the latter to the external structural constraints on head movements that we have described here.Video 1.Human cardiac thick filament reconstruction (EMD accession no. 2240) fitted with human cardiac atomic model (Protein Data Bank accession no. 5TBY) to show apparent absence of interaction between S2 from one level of heads and the SH3 and converter domains (red) of the next level. M-line at top. See also Fig. 3 B. Compare with Video 2.Video 2.Tarantula thick filament reconstruction (EMD accession no. 1950) fitted with tarantula IHM atomic model (Protein Data Bank accession no. 3JBH) to show 3-D view of interaction between S2 from one level of heads and the SH3 and converter domains (red) of the next level. M-line at top. See also Fig. 3 A and Fig. 4, A and B.Additional evidence comes from smooth muscle and nonmuscle myosin molecules, which can exist in a switched-off state in which the ATP turnover rate is similar to that in tarantula—also an HRX state (Cross et al., 1988). Strikingly, these myosins have a folded conformation, in which the two heads form an IHM, and the tail folds into three segments, the middle segment wrapping around the BH (Fig. 1 B; Suzuki et al., 1982; Trybus et al., 1982; Craig et al., 1983; Burgess et al., 2007; Yang et al., 2019). Cryo-EM analysis of this conformation shows intimate contact of the tail with the BH motor as it runs through the groove formed by the BH SH3 and converter domains (Fig. 4, C and D; Scarff et al., 2020; Yang et al., 2020). The regions of tail contact with the BH in these single molecules are very similar to those in the tarantula filament and would sterically impede BH converter movement, contributing to the HRX turnover rate of this inhibited form of the myosin molecule (Fig. 4). Although the inhibitory regions of the tail (S2 in tarantula, the middle segment of the tail in smooth muscle and nonmuscle myosin) and the nature of the interaction (intermolecular in one, intramolecular in the other) are both different, the likely inhibitory effects on BH converter movement and ATPase activity appear to be similar, supporting the importance of this interaction in generating the HRX state. HMM from these myosins lacks the distal two-thirds of the tail and therefore the interaction of the middle segment with the BH. Correspondingly, HMM exhibits an SRX but not an HRX rate (Cross et al., 1988). Together these observations provide strong support for the notion that interaction of the myosin tail with the SH3/converter region of the BH is the structural basis of hyper-relaxation.Another invertebrate muscle in which a putative HRX state has been observed is the scallop striated adductor. When thick filaments lose their ATP, helical ordering of the heads is lost (we would suggest by straightening of the heads in the apo state, so that the IHM can no longer form). EM shows that this process takes up to 30 min in scallop (Vibert and Craig, 1985), consistent with an HRX turnover rate. Scallop thick filaments differ from tarantula in having sevenfold rather than fourfold rotational symmetry (Vibert and Craig, 1983). Cryo-EM shows that this tightly packs the IHMs around the circumference of each crown, creating intermolecular interactions within crowns involving the SH3 domain of the BH and the motor domain of its neighbor FH (Figs. 1 B and 3 C; Woodhead et al., 2013; the heads in tarantula crowns are more widely spaced and do not show these interactions). While the reconstruction does not provide detail on possible tail interactions between crowns (as in tarantula), these additional intermolecular interactions may constrain the heads within a crown, impeding structural changes of the BH and FH and accounting for the hyper-slow release of products from scallop thick filaments through a mechanism quite different from that in tarantula.How is the IHM formed?Formation of the IHM depends on several key properties of myosin II: flexing of the heads and the head–tail junction and interaction of the motor domains with each other and with the proximal region of S2. BH–S2 interaction (where S2 runs over the bent BH) appears to be the primary binding interaction of the IHM (Alamo et al., 2016), as it is observed in smooth muscle HMM even when the FH is not bound to the BH (Burgess et al., 2007); FH–S2 interaction without interaction of the BH and BH–FH interaction without S2 are not observed. Interaction of S2 with the BH can only occur when the BH folds back onto the tail, and then only when the BH is in the strongly bent (Rosenfeld et al., 1994; Alamo et al., 2008, 2016; Scarff et al., 2020; Yang et al., 2020), nucleotide-trapping state, putatively with the SRX rate (Anderson et al., 2018). Thus, we picture the myosin molecule as having flexible heads (in a bent–straight equilibrium biased 90% toward the straight [DRX] conformation), which are flexibly attached to the tail, with the following sequence for formation of the IHM. If a transiently bent head comes in contact with S2, it binds and is stabilized in its bent (SRX) state, thereby becoming a BH (i.e., a precursor IHM; Alamo et al., 2016). The other head, flexing around its head–tail junction and in a similar bent–straight equilibrium, can now be caught (becoming an FH) when its converter region contacts the captured BH motor domain, stabilizing the FH bent conformation (Liu et al., 2003; Alamo et al., 2016; Scarff et al., 2020; Yang et al., 2020). This interaction is strengthened by contact of loops on the FH with S2 (Alamo et al., 2008; Scarff et al., 2020; Yang et al., 2020).Structural observations show that both heads of the IHM are strongly bent (Wendt et al., 2001; Woodhead et al., 2005; Scarff et al., 2020; Yang et al., 2020), while the bent structure in isolated heads may be uncommon. The sequence proposed above suggests how the IHM can be formed despite these constraints. If the bent structure occurs only rarely (e.g., 10% of the time), the likelihood of two bent heads coming together will be low: this likelihood is greatly increased by initial stabilization of one bent head (the BH) binding to S2 (forming the precursor IHM) and then binding of the second head (the FH) to the already bent BH of the precursor. This kinetic argument would explain why head–head interaction without involvement of the tail has not been observed, consistent with mutational studies (Adhikari et al., 2019). The final IHM is a tripartite structure with multiple weak interactions (BH–FH, BH–S2, and FH–S2) that inhibit ATPase activity as well as actin-binding capability (Scarff et al., 2020; Yang et al., 2020). These interactions are easily broken and in equilibrium with the noninteracting form. The result in isolated sarcomeric myosin soluble fragments at physiological ionic strength is an overall 25% SRX and 75% DRX rate (Anderson et al., 2018; Rohde et al., 2018; Gollapudi et al., 2021b). In filaments, individual IHMs (with the inhibited, bent structure of the individual heads) are stabilized by intermolecular interactions occurring in the polymer (as described earlier), increasing the fraction of SRX heads above 50%.Is there a relationship of the SRX state with animal lifestyle?We have suggested that nature uses two ways to enhance the SRX state: increase in fraction of inhibited heads and increase in degree of inhibition. The particular mechanism may be adapted to the lifestyle of the animal (Naber et al., 2011). Tarantulas spend long periods of time immobile and can survive many months without food. Minimizing ATP use during this time by hyper-relaxation of their BHs and SRX of most of their FHs would be an evolutionary advantage (Naber et al., 2011). They would nevertheless be ready for a rapid switch to the active state through the small numbers of swaying FHs that sense thin filament activation when muscle is stimulated (Brito et al., 2011). Scallops can swim quickly for short bursts by jet propulsion, using their striated adductor muscles to rapidly close their shell (e.g., to escape predators), but they remain stationary during extended periods of filter feeding (Speiser and Wilkens, 2016). In this low-activity state, the shell is held partially closed through contraction of the tonic smooth adductor muscle, which enters a catch state, maintaining force with little energy expenditure (Chantler, 2016). The striated adductor is 10 times more massive than its smooth counterpart (Naidu, 1987) and a potential metabolic drain during the long periods when the muscle is not in use: minimizing ATP use during these nonswimming periods through hyper-relaxation of their heads would be advantageous. The folded form of vertebrate smooth muscle and nonmuscle myosin is thought to serve as a storage molecule, which can be activated to form functional filaments as required through phosphorylation of their RLCs (Cross et al., 1988). The complete switching off of ATPase activity through hyper-relaxation when the molecule is not in use would again provide an evolutionary advantage.Vertebrate striated muscle thick filaments exhibit SRX but not hyper-relaxation. Interaction of IHMs with each other or with the filament backbone, MyBP-C, or titin enhances the fraction of heads in the SRX state but does not significantly increase the level of inhibition of the heads. SRX appears to be developed most strongly in the C-zone (probably in the two crowns of heads showing IHMs in each three-crown repeat), implicating MyBP-C in this inhibition (McNamara et al., 2016, 2019; Nelson et al., 2020). Myosin heads in the D-zone and in the non-IHM crown in the C-zone are less ordered (Zoghbi et al., 2008; AL-Khayat et al., 2013; Brunello et al., 2020) and may be the main source of DRX heads detected by single turnover experiments (Nelson et al., 2020). Vertebrates may make a trade-off of greater (though still low) ATP use in the RX state (SRX with some DRX heads) for the ability to switch on instantly as needed and for fine-tuning of the contractile response through interactions with other proteins such as MyBP-C.How are thick filaments activated from the SRX state?We have painted a picture in which most myosin heads in the thick filaments of resting muscle are tacked down on the filament surface in helices of IHMs in an SRX (or HRX) state. However, muscles must be ready for immediate activation from this energy-saving state. The other part of the picture therefore includes a small fraction of heads dissociated from the IHM at any particular moment. These relatively few, mobile, constitutively on (“sentinel”) heads are presumed to constantly explore the interfilament space, able to instantly sense thin filament activation and bind to actin when myosin-binding sites are exposed (by Ca²⁺-induced tropomyosin movement; Gordon et al., 2000), leading to initial tension development (Linari et al., 2015; Irving, 2017). We suggest that these are the transiently dissociated (swaying) FHs of the IHMs. In vertebrates, the sentinel heads could also include the less well-ordered heads at every third crown of the C-zone (Zoghbi et al., 2008; AL-Khayat et al., 2013) or the disordered heads in the D-zone (Brunello et al., 2020; Nelson et al., 2020). This essential role for a small number of mobile heads as the trigger for thick filament activity, when thin filaments are switched on, is paid for by the increase in ATP consumption over that used by SRX heads. What happens next depends on the type of muscle.In tarantula, the small amount of thick filament activity in an initial twitch appears to be enhanced in additional twitches by Ca²⁺-induced activation of myosin light chain kinase, which phosphorylates the FH RLCs, leading to increased release of the FHs from the IHMs, reduction in the HRX fraction and increase in the DRX fraction (Naber et al., 2011), and increased force production (Padrón et al., 2020). Prolonged high Ca²⁺ (e.g., in a tetanus) subsequently phosphorylates the BHs, resulting in their release and further enhancement of force (Padrón et al., 2020).Vertebrates have a finely tuned, graded response to activation. X-ray studies show that low-force isotonic contractions use only a small fraction of available heads (Linari et al., 2015). The majority remain helically ordered in their IHM configurations, continuing to save energy—even during contraction—representing a highly efficient use of ATP. As force increases, stress on the thick filaments rises, resulting in release of more heads, which produce greater force, in a positive feedback loop (Linari et al., 2015; Irving, 2017). In this mechanosensing model for thick filament activation, the filament is stretched by ∼1%, which may be sufficient to weaken intermolecular contacts between heads along the helical tracks that help to maintain the IHM conformations (Irving, 2017). Destabilization of the IHMs by stress could thus release the additional heads needed for strong force production. When contraction is switched off by removal of Ca²⁺ from the cytosol, IHMs rapidly go back to their helically ordered arrangement (Linari et al., 2015; cf. Ma et al., 2019), quickly returning the muscle to its energy-saving SRX state.In scallop, the thick filaments are directly activated by Ca²⁺ binding to the essential light chains (ELCs) on the myosin heads rather than through Ca²⁺ activation of the thin filaments (Szent-Györgyi et al., 1999; Chantler, 2016). Ca²⁺ binding causes breakage of the IHM intra- and intermolecular interactions, cooperatively releasing heads from their SRX/HRX state (Szent-Györgyi et al., 1999; Zhao and Craig, 2003; Jung et al., 2008; Chantler, 2016). Released Ca²⁺-activated heads can move freely, similarly to phosphorylated heads of tarantula, interacting with actin to generate force. As this process involves binding of Ca²⁺ to the myosin heads and not the slower, enzymatic phosphorylation of the light chains, full activation can occur rapidly (Zhao and Craig, 2003), making all heads immediately available for the powerful twitches that produce the strong swimming motions of this species. In the absence of a thin filament switch in scallops, sentinel heads may not be required as the heads directly sense Ca²⁺ activation.Interactions between IHMs along their helical tracks are common to all thick filaments in the SRX/HRX state. Reconstructions of tarantula, scallop, and vertebrate thick filaments, representing three distinct mechanisms of activation, all show such interactions, typically between the FH motor domain of one IHM and the BH regulatory domain of its neighbor in the next crown closer to the filament center (Figs. 2 C and ​and3;3; Woodhead et al., 2005; Zoghbi et al., 2008; AL-Khayat et al., 2013; Woodhead et al., 2013). As described earlier, these intermolecular interactions appear to stabilize the IHM conformation, enhancing the fraction of SRX heads in filaments. Concomitantly, they may also underlie rapid thick filament activation, as they provide a direct physical path for cooperative disruption of the IHM and thus exit from the SRX state (Stewart et al., 2010; McNamara et al., 2015). Cooperative exit from the SRX state has been experimentally demonstrated in skeletal and cardiac muscle incubated with mantATP chased by ADP (Stewart et al., 2010; Cooke, 2011; Gollapudi et al., 2021b). Cooperative activation is especially well developed in scallop thick filaments (Szent-Györgyi et al., 1999; Chantler, 2016), where it may underlie the rapid Ca²⁺ activation leading to the strong swimming motions of this species. The intermolecular interactions along helices and around each crown in scallops (Fig. 3 C; Woodhead et al., 2013) connect all heads in an extensive network that could be rapidly disrupted by Ca²⁺ binding.How is SRX modulated?The stability of the IHM and the level of SRX can be modulated in several ways, including myosin RLC phosphorylation and, in vertebrates, phosphorylation of MyBP-C (Stewart et al., 2010; Nag and Trivedi, 2021). RLC phosphorylation in tarantula greatly reduces the fraction of SRX and HRX heads and their ATP turnover times (Naber et al., 2011) while increasing DRX heads. This correlates with a decrease in helical ordering and extension of heads from the filament backbone (suggesting disruption of the IHM), as demonstrated by x-ray diffraction (Padrón et al., 2020) and EM (Craig et al., 1987). Phosphorylation not only activates heads but also maintains a memory of activation following the extensive phosphorylation that occurs in a tetanus; this can greatly potentiate subsequent contraction (Padrón et al., 2020). Disruption of the IHM and extension of heads toward neighboring actin filaments in the RX period following a tetanus presumably enables the stronger and more rapid interaction with actin of a post-tetanic contraction (Padrón et al., 2020). While the phosphorylated RX state that follows a tetanus would temporarily consume more ATP, it could provide a survival benefit by enabling stronger contractions when escaping predators or capturing prey. Following such periods of activity, the RLCs again become dephosphorylated, and the energy-saving SRX/HRX state, with ordered, interconnected IHMs lying along the filament surface, returns (Padrón et al., 2020), characterizing the long periods of inactivity in the life of the tarantula, when ATP savings are critical.Vertebrate skeletal muscle RLCs can also be phosphorylated, and phosphorylation correlates with post-tetanic potentiation (Sweeney et al., 1993) together with disordering of myosin helices and extension of heads from the filament surface (Levine et al., 1996; Yamaguchi et al., 2016), again suggestive of IHM disruption. As with tarantula, a corresponding reduction in the SRX state with phosphorylation (Stewart et al., 2010; Cooke, 2011; Gollapudi et al., 2021b), with a temporarily greater resting ATP consumption, pays for the greater contractility available following phosphorylation.Vertebrates have a second means of modulating the SRX state, which may provide finer control of thick filament activation/relaxation than with invertebrates. MyBP-C binds to myosin in the middle one-third of each half of the thick filament (the C-zone; Craig and Offer, 1976; Flashman et al., 2004; Luther et al., 2008), and several lines of evidence demonstrate that the SRX state is more pronounced in these regions (McNamara et al., 2016, 2019; Nelson et al., 2020; Nag and Trivedi, 2021), reaching as high as 90% (Nelson et al., 2020); this correlates with the clearest delineation of IHMs in reconstructions of the thick filament C-zone (Zoghbi et al., 2008; AL-Khayat et al., 2013). Toward the tips of the filament (the distal or D-zone), heads are less ordered (Brunello et al., 2020; R. Craig, unpublished EM data), and the SRX state is diminished (Nelson et al., 2020). In MyBP-C knockout mice, the IHM configuration is weakened or abolished (Zoghbi et al., 2008), and the SRX state is disrupted (McNamara et al., 2016). Thus MyBP-C appears to enhance the SRX state, apparently by stabilizing the IHM. This stabilization is further modulated by phosphorylation of MyBP-C, occurring in the heart in response to β-adrenergic stimulation. Enhancement of cardiac contractility by cardiac MyBP-C (cMyBP-C) phosphorylation may result in part from depression of cMyBP-C’s stabilizing effect on the SRX, which coincides with weakening of the IHM (Kensler et al., 2017; Caremani et al., 2019b; Irving and Craig, 2019; McNamara et al., 2019). These data overall imply that MyBP-C enhances energy saving by stabilizing the IHM structure and that this is modulated in the heart by cMyBP-C phosphorylation (McNamara et al., 2019).Importantly, in the healthy heart, RLC and MyBP-C phosphorylation are not zero but ∼50% (Chang et al., 2015) and ∼60% of maximum (Previs et al., 2012), respectively. This would suggest a partial weakening of the SRX state (compared with zero phosphorylation) during normal cardiac activity, which may poise myosin heads optimally between sequestration in the IHM (to save energy) and availability for interaction with actin to generate force, with fine-tuning of these levels available upon further phosphorylation. Thus, we assume that the level of SRX of a muscle (degree of IHM formation) is tuned to the physiological needs of the moment, with the goal over time of minimizing energy consumption within these limits. Strikingly, phosphorylation levels of MyBP-C and RLC can both decrease in heart failure (El-Armouche et al., 2007; Toepfer et al., 2013), which would predict a higher fraction of SRX heads. This may reduce the need for energy under these adverse circumstances but may also contribute to the compromised contractility of the failing heart.Animals can save energy when food supplies are scarce or weather conditions adverse by a reduction in body temperature. This occurs naturally in ectotherms (cold-blooded animals) when exterior temperatures drop and by hibernation in some endotherms (warm-blooded animals), where body temperature and metabolism reduce to low values. Does enhanced SRX play a role in energy conservation under these circumstances? X-ray diffraction of both mammalian and tarantula muscle at low temperatures (10°C) suggests that the number of myosin motors in the helically ordered, IHM conformation decreases substantially compared with temperatures nearer physiological levels (Malinchik et al., 1997; Xu et al., 1997; Caremani et al., 2019a, 2021; Ma et al., 2021a); modeling of tarantula suggests that it is specifically the FHs that are disordered, while the BHs remain ordered (Ma et al., 2021a). This disordering suggests that the SRX state may actually be reduced, rather than enhanced, in cold temperatures. Other factors may contribute to energy conservation by muscle under cold conditions (Caremani et al., 2021; Ma et al., 2021a): myosin heads become refractory to actin binding at low temperature (Caremani et al., 2019a), and the disordered FHs, containing ADP.Pi at the active site, may transition toward the ATP conformation, inhibiting ATP turnover (Xu et al., 1999; Ma et al., 2021a). The impact of torpor (a form of hibernation) on the SRX state has recently been explicitly studied in the 13-lined ground squirrel (Ictidomys tridecemlineatus; Toepfer et al., 2020). During torpor, core body temperature drops to 5°C, metabolic rate to 3% of basal levels, and heart rate from 311 to 6 beats per min; hummingbirds undergo a similar dramatic reduction in metabolic rate to save energy each night (Shankar et al., 2020). The fraction of SRX heads found in cardiac muscle removed from the ground squirrel during torpor was reported to increase from 65 to 75%, contrary to expectation from the low-temperature x-ray studies of mammalian muscle described above that would imply a decrease in the number of IHMs. This discrepancy could be due to performance of the SRX measurements at 21°C (Toepfer et al., 2020) rather than the low temperatures used in the x-ray experiments that revealed helical disordering. The latter would presumably reflect the thick filament structure most accurately at actual torpor temperatures. We speculate that the phenomenon of iguanas (ectotherms) falling from trees and becoming immobile when environmental temperatures are abnormally low (Stroud et al., 2020) may be caused in part by the refractory impact of temperature on the ability of their myosin heads to bind to actin; a similar effect may have contributed to extinction of the dinosaurs during the global cooling that followed the asteroid impact of 66 million years ago (Vellekoop et al., 2014).Modulation of the SRX and its putative structural correlate, the IHM, may play a critical role in a number of cardiac diseases and their treatment, summarized elsewhere (Alamo et al., 2017; Nag et al., 2017; Yotti et al., 2019; Trivedi et al., 2020; Daniels et al., 2021; Schmid and Toepfer, 2021). Hypertrophic cardiomyopathy (HCM) is an inherited disease caused by mutations in sarcomeric proteins, including myosin, and characterized by hypercontractility and the inability to fully relax during diastole (Ashrafian et al., 2011). Recent studies show that mutations in the myosin heavy chain (MYH7; accounting for ∼40% of HCM cases) strongly cluster in regions of the myosin molecule involved in the interfaces of the IHM (Alamo et al., 2017; Nag et al., 2017), and lead to a substantial decrease in SRX and increased energy use (Anderson et al., 2018; Adhikari et al., 2019; Sarkar et al., 2020). Disruption of the IHM by such mutations, at the head–head or BH–S2 interface, could release more heads for interaction with actin, accounting for the hypercontractility and impaired relaxation observed (Alamo et al., 2017; Anderson et al., 2018; Spudich, 2019; Sarkar et al., 2020). Experimental evidence for this proposal has been obtained (Adhikari et al., 2019; Sarkar et al., 2020). Mutations in MyBP-C leading to HCM also appear to partially disrupt the SRX state of the myosin heads (Toepfer et al., 2019; Nag and Trivedi, 2021), leading to an increase in the number of DRX heads, which may contribute to the observed hypercontractility (McNamara et al., 2017). Depending on the mutation, cMyBP-C may have a reduced affinity for myosin, or the amount of MyBP-C incorporated into the thick filament may be reduced. It has been proposed that either mechanism would reduce the strength of the cMyBP-C–myosin interaction, releasing heads from the filament backbone (DRX heads), which could be partially responsible for the hypercontractile phenotype observed in patients with these mutations (Colson et al., 2007; Toepfer et al., 2019; Nag and Trivedi, 2021).Impaired relaxation due to HCM mutations in myosin (and other myofibrillar proteins) can be compensated by a recently developed drug, mavacamten, which has been shown to increase the fraction of myosin heads in the SRX state (Anderson et al., 2018; Rohde et al., 2018; Spudich, 2019; Nag and Trivedi, 2021), specifically in the D-zone of thick filaments (Nelson et al., 2020), concomitant with an increased number of molecules folded into the IHM conformation (Anderson et al., 2018; Rohde et al., 2018; Gollapudi et al., 2021b). In thick filaments of intact cardiac muscle, mavacamten increases the degree of quasi-helical ordering of myosin heads, consistent with an increase in the stability or fraction of molecules in the IHM conformation (Anderson et al., 2018), supporting our overall contention that the IHM is the main basis of the SRX state in muscle. Mavacamten inhibits the ATPase activity of isolated S1, specifically by inhibiting phosphate release (Green et al., 2016), suggesting that it enhances the bent (SRX) state of the myosin head, which, based on our earlier reasoning, would lead to the observed increase in the fraction of molecules in the IHM (Anderson et al., 2018).Future directionsWe currently know the structure of the IHM at ∼15 Å resolution in tarantula (Yang et al., 2016) and insect (Hu et al., 2016) filaments and ∼4 Å resolution in single smooth muscle myosin molecules (Scarff et al., 2020; Yang et al., 2020). Improvements in resolution should enhance visualization of the side-chain interactions that stabilize the SRX state in both filament and monomer. For filaments, cryo-EM of tarantula offers the greatest potential, owing to the stability of its helices. To better understand how mutations in the IHM interfaces lead to the hypercontractility of HCM, cryo-EM of vertebrate cardiac thick filaments is required to improve the resolution beyond the current ∼40 Å (obtained with negative staining; Zoghbi et al., 2008; AL-Khayat et al., 2013), a difficult task owing to the lability and quasi-helical symmetry of the vertebrate thick filament head array. Differences in the levels of SRX in the C- and D-zones of the thick filaments (Nelson et al., 2020) suggest differences in IHM stability or interactions, which will need to be analyzed by cryo-EM, again a significant task for the reasons stated above and because of the small size of these zones. The most detailed insights into IHM intramolecular interactions should come from a high-resolution structure of the IHM in isolated cardiac myosin molecules, which is urgently needed. This could potentially be obtained by x-ray crystallography or cryo-EM of IHM constructs (e.g., 25 heptad; Fig. 1; Anderson et al., 2018), although this is likely to be hampered by the relative instability of the structure. Incubation with mavacamten may improve its stability (Anderson et al., 2018). How interaction of MyBP-C with myosin enhances the SRX state is another fertile but challenging area of investigation, which may require cryo-electron tomography of thick filaments or intact myofibrils (Burbaum et al., 2021) or single particle cryo-EM studies of MyBP-C–IHM complexes. Experiments suggest that thick filaments in muscle are in equilibrium with a pool of myosin monomer, which may play a role in thick filament assembly/disassembly during development, hypertrophy, and myosin turnover (Saad et al., 1986; Katoh et al., 1998; Ojima, 2019). Myosin monomers at physiological ionic strength form IHMs with a folded tail structure and slow ATP turnover rate, which may act as a transport form from ribosome to filament (Ankrett et al., 1991; Katoh et al., 1998; Jung et al., 2008). Whether the SRX/IHM structure in thick filaments affects the monomer–polymer equilibrium is an area for future investigation.ConclusionThe SRX state of thick filaments plays a crucial role in the energy balance of muscle and is ubiquitous across the animal kingdom. It results from a conformation of the myosin head in which ATP turnover is strongly inhibited, minimizing ATP use. This conformation is stabilized by intramolecular interactions when it is incorporated into the IHM and by additional intermolecular interactions when IHMs are assembled into thick filaments, increasing the fraction of energy-saving SRX heads. Thus, we propose that IHMs, helically organized along the thick filament surface in the relaxed state, are the major basis of SRX in living muscle. An HRX state, found in thick filaments of some invertebrates and in the folded (storage) form of smooth muscle and nonmuscle myosin molecules, is also based on the IHM but in these cases results from additional intra/intermolecular interactions. A filament in which every head was in SRX, while maximally saving ATP, would not be useful in contraction: at any time, a small fraction of heads is dissociated (DRX or sentinel heads) and available to sense thin filament activation and generate initial force. The SRX state is down-regulated in situ by RLC and MyBP-C phosphorylation in response to physiological requirements (activation), which disrupt the IHM, releasing heads for interaction with actin. Mutations in myosin or MyBP-C causing HCM disrupt the SRX state, causing hypercontractility, which can be reversed by drugs that stabilize the IHM and thus the SRX.Online supplemental materialVideo 1 shows human cardiac thick filament reconstruction fitted with a human cardiac atomic model to demonstrate an apparent absence of interaction between S2 from one level of heads and the SH3 and converter domains of the next level. Video 2 shows tarantula thick filament reconstruction fitted with a tarantula IHM atomic model to show that in this species there is an interaction between S2 from one level of heads and the SH3 and converter domains of the next level.     

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.