Structural Basis of Tubulin Recruitment and Assembly by Tumor Overexpressed Gene (TOG) domain array Microtubule Polymerases

XMAP215/Stu2/Alp14 proteins accelerate microtubule plus-end polymerization by recruiting tubulins via arrays of Tumor Overexpressed Gene (TOG) domains. The underlying mechanism of these arrays as microtubule polymerases remains unknown. Here, we describe the biochemical and structural basis for TOG domain arrays in recruiting and polymerizing tubulins. Alp14 binds four tubulins via dimeric TOG1-TOG2 arrays, each with distinct exchange rates. X-ray structures reveal pseudo-dimeric square-shaped assemblies in which four TOG domains position four unpolymerized tubulins in a polarized wheel-like configuration. Crosslinking confirms square assemblies form in solution, and inactivation of their interfaces destabilizes square organizations without influencing tubulin binding. Using an approach to modulate tubulin polymerization, we determined a X-ray structure showing an unfurled assembly in which TOG1 and TOG2 uniquely bind two polymerized tubulins. Our findings suggest a new microtubule polymerase model in which TOG arrays recruit tubulins by forming square assemblies, which then unfurl facilitating their concerted polymerization into protofilaments.


Introduction:
Microtubules (MTs) are highly dynamic polarized polymers that perform critical cellular functions including forming bipolar mitotic spindles, intracellular organization, and modulating cell development and cell migration Steinmetz, 2008, 2015). MTs are assembled from αβ-tubulin heterodimers (αβ-tubulin) and their polymerization exhibits dynamic instability arising from guanosine triphosphate (GTP) hydrolysis in β-tubulins at MT ends. However, the conformational changes promoting soluble αβ-tubulins to polymerize at MT ends remain poorly understood. The polymerization of αβ-tubulin and its GTP hydrolysis are regulated by conserved proteins that bind MT plus-ends or along MT lattices (Akhmanova and Steinmetz, 2008Al-Bassam and Chang, 2011;Al-Bassam et al., 2010;Brouhard and Rice, 2014). The XMAP215/Stu2/Alp14 family of MT polymerases is among the best-studied MT regulators. They localize at the extreme tips of MT plus-ends and accelerate αβtubulin polymerization in eukaryotes Steinmetz, 2011, 2015;Al-Bassam and Chang, 2011;Maurer et al., 2014). Loss or depletion of MT polymerases is lethal in several organisms as it severely decreases MT polymerization rates during interphase resulting in shortened mitotic spindles in most eukaryotes studied (Al-Bassam et al., 2012;Cullen et al., 1999;Wang and Huffaker, 1997). MT polymerases also bind kinetochores where they accelerate MT dynamics and regulate kinetochore-MT attachment (Miller et al., 2016;Tanaka et al., 2005). MT polymerases recruit αβ-tubulins via an array of conserved Tumor Overexpressed Gene (TOG) domains (termed TOG arrays, from herein), which are critical for their function (Reber et al., 2013;Widlund et al., 2011). Arrays of TOG-like domains are conserved in two other classes of MT regulators, CLASP and Crescerin/CHE-12 protein families (Al-Bassam and Chang, 2011;Al-Bassam et al., 2010;Das et al., 2015), suggesting that arrays of TOG domains uniquely evolved to regulate diverse MT polymerization functions through binding αβ-tubulins in different intracellular settings.
Yeast MT polymerases, such as S. cerevisiae Stu2p and S. pombe Alp14, are homodimers consisting of two unique and consecutive TOG domains, TOG1 and TOG2, per subunit numbered based on their location from the N-terminus. In contrast, metazoan orthologs, such as XMAP215 and ch-TOG, are monomers including five tandem TOG domains, TOG1 through TOG5, (Al-Bassam and Chang, 2011;Brouhard and Rice, 2014). Phylogenetic analyses suggest that TOG1 and TOG2 domains are evolutionarily distinct (Al-Bassam and Chang, 2011), and the TOG3 and TOG4 domains in metazoans are evolutionarily and structurally related to the TOG1 and TOG2 domains, respectively (Brouhard et al., 2008;Fox et al., 2014;Howard et al., 2015). Thus, despite differences in TOG array organization in yeast and metazoan proteins, both groups contain at least two sets of tandem TOG1-TOG2 domains.
Structural studies contribute to our understanding of the molecular basis of TOG domain function in recruiting soluble αβ-tubulin. Each TOG domain is composed of six α-helical HEAT (Huntingtin, EF3A, ATM, and TOR) repeats, which forms a conserved paddle-shaped structure (Al-Bassam and Chang, 2011;Al-Bassam et al., 2007;Al-Bassam et al., 2006;Brouhard and Rice, 2014;Slep and Vale, 2007). X-ray structures of isolated TOG1 and TOG2 domains in complex with αβtubulins reveal that these domains recognize the curved αβ-tubulin conformations via inter-helical loops positioned along an edge of these paddleshaped domains (Ayaz et al., 2014;Ayaz et al., 2012). Straightening of the curved soluble αβ-tubulins upon polymerization into MTs likely dissociates TOG domains. Our previous studies indicate that native TOG arrays from yeast or metazoan MT polymerases assemble into discrete particles upon binding αβtubulin (Al-Bassam and Chang, 2011;Al-Bassam et al., 2006;Brouhard and Rice, 2014). Both TOG1 and TOG2 domains are critical for MT polymerase function and their inactivation in fission or budding yeast MT polymerases results in MT function defects (Al-Bassam et al., 2012;Al-Bassam et al., 2006;Ayaz et al., 2014). Two models were proposed to explain how arrays of TOG domains function as MT polymerases: One model, which is based on studies of native TOG arrays, indicates that TOG arrays may form ordered assemblies upon binding αβ-tubulins (Al-Bassam et al., 2006;Brouhard et al., 2008). A second model, based on studies of isolated TOG domains or short TOG arrays suggests these arrays form flexible assemblies in which TOG1 and TOG2 independently recruit multiple αβ-tubulins to MT plus-ends (Al-Bassam and Chang, 2011;Ayaz et al., 2014). Distinguishing between these models requires understanding the high-resolution organization of native TOG arrays in complex with αβ-tubulin and their transitions during the αβ-tubulin recruitment and polymerization phases.
Here, we describe biochemical and structural analyses of TOG arrays during αβtubulin recruitment and polymerization states. We show that dimeric yeast MT polymerases recruit αβ-tubulins using TOG1 and TOG2 domains, which bind and release αβ-tubulins with unique exchange rates. X-ray structures of αβ-tubulin-TOG array complexes reveal pseudo-dimeric TOG1-TOG2 subunits form a headto-tail square-shaped assembly, which orients αβ-tubulins in a polarized configuration. Crosslinking and mass spectrometry show dimeric yeast TOG arrays that form square conformations, confirming that these states exist in solution. Mutants in which these interfaces are inactivated show disrupted square organization, without defects in αβ-tubulin binding. Using a novel approach to promote the limited polymerization of αβ-tubulin while bound to TOG arrays, we determined a x-ray structure of an "unfurled" assembly revealing TOG1-TOG2 domains are bound onto two αβ-tubulins polymerized head-to-tail into a protofilament. These studies establish a new "polarized unfurling" model for TOG arrays as MT polymerases. In the accompanying manuscript, we present in vitro reconstitution studies and in vivo functional studies in the fission yeast S. pombe using two classes of Alp14 structure-based mutants, which provide evidence supporting various facets of this new model.

Results
Arrays of TOG1 and TOG2 domains recruit multiple αβ-tubulins via and exhibit unique affinities and exchange rates.
First, we studied the binding affinities and stoichiometries of near native monomeric and dimeric yeast TOG arrays to bind αβ-tubulin. A yeast TOG array consists of TOG1 and TOG2 domains linked by 55 to 80-residue linker and can be dimerized via a C-terminal coiled-coil. Individual TOG domains bind αβtubulins via a narrow interface, which involves mostly ionic contacts (Ayaz et al., 2014;Ayaz et al., 2012). Thus, we explored how changes in ionic conditions influence αβ-tubulin binding stoichiometry of monomeric S. pombe Alp14 TOG1-TOG2 array (termed Alp14-monomer from herein, and includes residues 1-510) or dimeric Alp14 TOG array (termed wt-Alp14-dimer from herein: residues 1-690), using size-exclusion chromatography (SEC) and multi-angle light scattering (SEC-MALS) ( Figure 1A µM wt-Alp14-dimer bound to four αβ-tubulins via its four TOG domains, whereas the 1 µM wt-Alp14-monomer only bound to two αβ-tubulins via two TOG domains, suggesting TOG1 and TOG2 independently recruit αβ-tubulins ( Figure   1A Table   S1-S2). At 200 mM KCl conditions, however, both 1 µM wt-Alp14-monomer and wt-Alp14-dimer bound roughly half the expected αβ-tubulin compared to that at 80 or 100 mM KCl, resulting in lower-mass complexes. The latter suggests that either TOG1 or TOG2 domain, but not both, maintains binding to αβ-tubulin at 200 mM KCl conditions ( Figure 1A Table S1-S2); the values for binding stoichiometry at these 100 or 200 mM ionic conditions resolve the discrepancies between our previously reported binding stoichiometries and those reported by other groups (Al-Bassam et al., 2012;Al-Bassam et al., 2006). To determine if either TOG1 or TOG2 retains αβ-tubulin binding at 200 mM KCl, we studied Alp14 mutants in which TOG1 or TOG2 αβ-tubulin binding interfaces were inactivated through mutating conserved intra-HEAT repeat turn residues (see materials and methods; Figure Table S1-S2). Moreover, the αβtubulin binding molar ratios and the measured stoichiometry of TOG1M and TOG2M at 80-100 mM KCl, determined by SEC and SEC-MALS, respectively, were roughly half those measured for wt-Alp14-dimer, supporting the independent, and unique activities of TOG1 and TOG2 in recruiting αβ-tubulins ( Figure 1E).
We, next, quantitatively measured the absolute TOG1 and TOG2 affinities for αβtubulin and how those change due to changes in ionic strength (100-200 mM KCl) using isothermal titration calorimetery (ITC). ITC data show isolated TOG1 (residues 1-270) and TOG2 (residues 320-510) bind αβ-tubulins with roughly 2.5fold difference in dissociation constants ( Figure 1G; Figure 1-Supplement 3). At 100 mM KCl, we measured dissociation constants for TOG1 and TOG2 to be 70 and 173 nM, respectively. These data suggest both TOG1 and TOG2 exhibit a fairly high affinity for αβ-tubulin with 2.5-fold affinity difference and are nearly identical to those previously reported (Ayaz et al., 2014;Ayaz et al., 2012).
However, at 200 mM KCl, we measured TOG1 and TOG2 αβ-tubulin dissociation constants at 1.5 µM and 3.2 µM, respectively, which are 20-fold lower in absolute affinity compared to those measured at 100 mM KCl. Together, our studies suggest that at 100 mM KCl or below, each TOG1-TOG2 array subunit tightly binds two αβ-tubulins, whereas at 200 mM KCl it binds one αβ-tubulin tightly via TOG1 and exchanges a second αβ-tubulin rapidly via TOG2 ( Figure 1H). In the accompanying manuscript, we describe the implications of this ionic strength change in modulating the MT polymerase activity through influencing TOG2 (Cook et al., 2018). We also present in vitro and in vivo studies for TOG1M and TOG2M, suggesting that TOG1 and TOG2 domains, with their unique exchange rates for αβ-tubulin serve unique and non-additive roles in MT-plus end tracking and MT polymerase activity, respectively (Cook et al., 2018).

tubulins.
Our biochemical studies suggest that structural studies of TOG arrays:αβ-tubulin complexes must be conducted at 80-100 mM KCl and at high concentrations to avoid αβ-tubulin dissociation from TOG2 domains. To increase complex homogeneity and inhibit αβ-tubulin self-assembly under such conditions, we utilized designer ankyrin repeat protein, Darpin-D1 (termed DRP from herein), which specifically binds β-tubulin's polymer-forming interface (Pecqueur et al., 2012). We studied if DRP binding to αβ-tubulin influences the ability of wt-Alp14monomer or wt-Alp14-dimer to bind αβ-tubulins. Binding molar ratios and stoichiometries were measured for DRP bound-αβ-tubulin wt-Alp14 complexes by SEC and SEC-MALS at 80-100 mM KCl conditions ( Figure 1E; Figure 1 Supplement 2A-G; Table S1-S2). These studies suggest that DRP does not affect the simultaneous binding of multiple αβ-tubulins to TOG arrays in monomer or dimer forms. At 1 µM concentration, wt-Alp14-dimer formed a complex with four αβ-tubulin and four DRP at a molar ratio of 2:4:4 ( Figure 1E;   Table S1). These ability of these tubulin to bind stoichiometric amounts of DRP suggest that αβ-tubulins recruited by TOG array are in a non-polymerized state upon their initial association. This feature is consistent with a reported lack of cooperativity described between TOG1 and TOG2 in binding to αβ-tubulins (Ayaz et al., 2014). Accordingly, we used this strategy to identify crystallization conditions using yeast MT polymerase orthologs from a variety of organisms (see materials and methods).
Crystals of the Saccharomyces kluyveri ortholog of Alp14 (termed sk-Alp14monomer; residues 1-550) bound to αβ-tubulins and DRP grew in conditions similar to those used for SEC and SEC-MALS (Figure 2 Supplement 1A). Using crystals with either a native sk-Alp14-monomer or a sk-Alp14-monomer with a modified TOG1-TOG2 linker sequence, termed sk-Alp14-monomer-SL (see materials and methods; Figure 2 Supplement 2), we determined X-ray structures for 1:2:2 sk-Alp14 TOG1-TOG2: αβ-tubulin: DRP from complexes using sk-Alp14-monomer and sk-Alp14-monomer-SL with molecular replacement (see materials and methods) at 4.4 and 3.6-Å resolution, respectively (Table S3 and Supplement 2D-F). We anticipate sk-Alp14-monomer formed dimeric organization, despite a lack of dimerization domains, due to their high concentration during crystallization. The X-ray structures revealed two TOG1-TOG2 subunits in a pseudo-dimeric assembly forming the core of these complexes. In a TOG square, each TOG domain is bound to a curved αβ-tubulin capped by a DRP through its outward-facing binding interface, and is minimally contacted by the neighboring TOG-bound αβ-tubulin ( Figure 2A; Figure 2 Supplement 1F-H). The distances and interaction patterns between residues of α-tubulin and DRP bound onto a neighboring β-tubulin indicate that DRP only interacts with its cognate β-tubulin does not bind a neighboring α-tubulin ( Figure   2 Supplement 2H-K). The latter suggests DRP has no effect on stabilizing each TOG square assembly. DRP binding rather only caps β-tubulin, presenting a significant impediment to the polymerization of αβ-tubulin while bound to the TOG array. The αβ-tubulins bound onto the TOG square are positioned in a polarized orientation, by virtue of the asymmetry in the TOG domain αβ-tubulin interface and pseudo-dimeric TOG1-TOG2 subunit interfaces within the TOG square (see below). The β-tubulin on a TOG1 bound αβ-tubulin is rotated roughly 90° from its polymer-forming interface relative to the adjacent α-tubulin on a TOG2 bound αβ-tubulin ( Figure 2B).

Two interfaces stabilize pseudo-dimeric TOG array into a TOG square.
The X-ray structures reveal each TOG square is a dimer of TOG1-TOG2 array subunits assembled head-to-tail from alternating TOG1 and TOG2 domains.
TOG domains are aligned along their narrow edges, analogous to four links attached head-to-tail forming an asymmetric square-like complex with two edges slightly longer than their orthogonal edges ( Figure 2C, D). Two contact sites, which we term interfaces 1 and 2, stabilize the TOG square. These interfaces are formed by interactions formed via inter-HEAT repeat loops of each TOG domain, which are located on the opposite edges from the αβ-tubulin-binding sites.
Although TOG1 and TOG2 domains are each 60-Å long, the TOG square assembly is slightly rectangular with 115-Å by 98-Å dimensions due to wider overlaps between TOG1 and TOG2 domains leading to 10-Å stagger at interface 1 sites, in contrast to a direct end-on corner-like interface 2 sites. Both interfaces 1 and 2 are stabilized by hydrophobic packing and ionic interaction zones ( Figure   2E-H). Interface 1 packs a 668-Å 2 surface area, and positions the TOG1 Cterminus at 90° to a 10-residue segment of the TOG1-TOG2 linker and the Nterminus of TOG2. The TOG1-TOG2 linker sequence forms an extended polypeptide that critically bridges interactions between TOG1 HEAT repeat 6 αhelix/inter-HEAT 5-6 loop segment and the TOG2 inter-HEAT repeat 1-2/2-3 loop segments ( Figure 2E, F). Interface 2 packs a 290-Å 2 surface area and positions the TOG2 C-terminus at 90° to the N-terminus of TOG1 ( Figure 2G, H) In interface 2, the TOG2 inter-HEAT repeat 4-5 loop interacts with the TOG1 inter-HEAT repeat 1-2/HEAT1 α--helix ( Figure 2G). The residues forming interface 1 and interface 2 within TOG1, TOG2 and linker regions are either moderately or highly conserved ( Figure 2F, H; Figure 2 Supplement 2). The total buried surface area stabilizing two sets of interfaces 1 and 2 in a TOG square is 1930 Å 2 , which is moderate in size and dispersed for such a large assembly. This suggests that this conformation maybe meta-stable and that DRP binding and its inhibition of αβ-tubulin polymerization may have stabilized this intermediate. `

Dimeric Alp14 TOG arrays forms a TOG square conformation in solution
Next we examined and chemically trapped the direct physical interactions between TOG1 and TOG2 based on the interfaces observed in TOG square structure using cysteine crosslinking. We generated mutants with specific cysteine pairs within the two sides of interface 1 (S180C, L304C) or interface 2 (S41C and E518C) in native dimeric sk-Alp14 (termed sk-Alp14-dimer; residues 1-724) ( Figure 3A, B). We tested whether these interfaces form inter-subunit contacts in dimeric TOG array by crosslinking via disulfide oxidation. A 110-kDa . We also observed a ~170-kDa species which specifically formed in the sk-Alp14 S180C-L304C mutant, and not in the native sk-Alp14-dimer or the sk-Alp14-S41C-E518C mutants. Furthermore, this 170-kDa intermediate was also not observed with sk-Alp14-S180C-L304C without αβ-tubulin ( Figure 3D). Massspectrometry (LC/MS-MS) confirmed this 170-kDa intermediate is indeed the sk-Alp14-S180C-L304C protein. Next, we mapped the cysteine residues involved disulfides in sk-Alp14-S180C-L304C mutant through peptide disulfide mapping after differential alkylation and mass-spectrometry (see materials and methods).
This approach revealed only two classes of peptides in sk-Alp14 S180C-L304C with 105-Da mass added onto cysteines, suggesting they were engaged in disulfides, with the following sequence boundaries: 297-320 and 179-189 ( Figure   3 Supplement 1B). These two peptide sequences represent TOG1 inter-HEAT-repeat and TOG1-TOG2 linker regions both of which are involved in forming interface 1 in the x-ray structures ( Figure 2). All the remaining peptides with cysteine residues that were identified in sk-Alp14 S180C-L304C included 57-Da in added mass, suggesting they were in the reduced form and did not form disulfides. Thus, these data directly provide independent support that interface 1 of the TOG square conformation forms in sk-Alp14-dimer in solution outside the crystallographic setting, and is indeed the inter-subunit dimeric interface between two TOG arrays subunits, as visualized in the crystal structures ( Figure 3A, B).

Inactivation of interfaces 1 and 2 disrupts TOG square assembly but does not disrupt αβ-tubulin binding.
We explored the role of Interfaces 1 and 2 in formation of the TOG square assembly and the binding of TOG arrays to αβ-tubulins, We generated three Alp14-dimer mutants which harbor either partially or fully disrupted interfaces 1 and 2 sites ( Figure 4A). We targeted disruption of salt bridges or hydrophobic zones in interfaces 1 and 2 by mutating conserved alanines, leucines or glutamates ( Figure 4A). Charged residues were either replaced with alanines or residues of the opposite charge, and hydrophobic residues were replaced with charged residues to dissociate hydrophobic interactions. We mutated eight residues to disrupt interface 1 (termed INT1), seven residues to disrupt interface 2 (termed INT2) or fifteen residues to disrupt both interfaces 1 and 2 (termed .

Unfurling of TOG arrays promote concerted αβ-tubulin polymerization into protofilaments.
The TOG square conformation shows how αβ-tubulins are recruited but does not reveal how TOG arrays drive αβ-tubulin polymerization. We hypothesized that the TOG square structure may undergo a subsequent conformational change to promote polymerization of the recruited αβ-tubulins. To explore this transition, we created a biochemical approach to partially release αβ-tubulin polymerization by relieving the DRP inhibition. We reasoned that a structural transition may occur more readily if the DRP dissociates from β-tubulin in a crystallization setting. We developed a strategy to conditionally release αβ-tubulin polymerization while recruited into TOG arrays by using a weakened affinity DRP. We reasoned that the increased dissociation of DRP may allow complexes to form polymerized αβtubulin intermediates in steady state. To accomplish this, we removed the Nterminal ankyrin repeat of DRP (termed DRPΔN from herein). We measured DRPΔN affinity using ITC revealing a three-fold decrease in its αβ-tubulin binding affinity as compared to DRP ( Figure 5A (Table S3). These crystals exclusively formed only when DRPΔN was used with αβ-tubulin:sk-Alp14-monomer or dimer complexes. We determined an X-ray structure to 3.3-Å resolution by molecular replacement using these crystals ( Supplement 2C, D). TOG1 and TOG2 are specifically bound to the lower and upper αβ-tubulins, respectively, of a highly curved protofilament. Only a single DRPΔN caps the TOG2 bound αβ-tubulin ( Figure 5E). Compared to the TOG square, this "unfurling" rearrangement requires 68°-rotation and 32-Å translation of TOG2:αβ-tubulin hinging around interface 2 and TOG1-αβ-tubulin ( Figure 6A, B). This transition promotes the concerted polymerization of TOG2:αβ-tubulin onto the plus-end of TOG1:αβ-tubulin, and consequently driving the dissociation of the second DRPΔN ( Figure 5C, D). The two αβ-tubulin polymer in this complex is a highly curved protofilament (16.4° inter-dimer interface), and it displays ~3° more curvature than RB3/stathmin-αβ-tubulin curved protofilament structure (Table S5; Figure 5 Supplement 2E) (Nawrotek et al., 2011). Comparison of αβtubulin dimer structure (α2β2) within this 1:2:1 structure to the unpolymerized αβtubulin structure revealed polymerization is associated with a 5° rotation and 10-Å translation in the T7 loop and H8 helix in the TOG2-bound α-tubulin, which engages TOG1-bound β-tubulin elements and the E-site bound GDP nucleotide ( Figure 6C, D). The latter conformational change likely stabilizes inter-dimer αβtubulin interfaces ( Figure 5D), burying a 1650-Å inter-dimer interface ( Figure 5E, F). This conformational change occurs at similar site to those observed during the MT lattice GTP hydrolysis transition (Alushin et al., 2014). Thus the 1:2:1 unfurled structure represents a concerted αβ-tubulin post-polymerization intermediate promoted by a TOG1-TOG2 subunit prior to straightening the protofilament.

MT protofilament plus-ends.
We next evaluated how x-ray structures for αβ-tubulin loaded TOG square and unfurled assemblies may dock onto protofilament tips at MT-plus ends. Atomic models of for these states were overlaid onto the terminal αβ-tubulins of the curved GTP or GDP tubulin protofilament models (Figure 7). The four αβ-tubulin bound TOG square assembly x-ray structure (Figure 2) was superimposed onto the terminal αβ-tubulin at protofilament ends in two docking orientations, either via the αβ-tubulins bound onto TOG1 or TOG2 ( Figure 7A, B). We observe mild steric surface overlap between the four αβ-tubulin loaded TOG square and the protofilament when TOG1-αβ-tubulin is docked onto the protofilament end. This steric overlap caused by overlap between αβ-tubulin-TOG2 from the second TOG square subunit and the penultimate αβ-tubulin from the protofilament end ( Figure 7A; Figure 7 Supplement 1A). In contrast, we observe no steric contact when the TOG square is docked via αβ-tubulin-TOG2. In this orientation, the TOG1-αβ-tubulin from the second subunit is retracted by 10-Å from the penultimate αβ-tubulin in the protofilament ( Figure 7B). The differences between steric overlap between the TOG square and the protofilament in these two docking orientations are due to slightly asymmetric length and width dimensions of the TOG square, which are caused by stagger at interface 1. These differences suggest that the destabilization of the TOG squares is more likely if TOG1-αβ-tubulin docks onto the protofilament plus end, in contrast to TOG2-αβtubulin docking. The slow exchange rate of TOG1-αβ-tubulin makes this more likely while the rapid exchange of αβ-tubulin by TOG2 makes this less likely. The unfurled 1:2 TOG1-TOG2:αβ-tubulin assembly can only docked using TOG1-αβtubulin onto the protofilament plus-end, and suggests that TOG2:αβ-tubulin is positioned the furthest away from the MT-plus-end in this conformation. These models were used to assemble steps for a new MT polymerase model described in the discussion (Figure 8).

A "Polarized Unfurling" model for TOG arrays as MT polymerases
The combination of structural, biochemical analyses suggest a new model for TOG arrays as MT polymerases. We propose a "polarized-unfurling" MT polymerase model, as summarized in Figure 8 and animated in Video S1, and is supported by docking models for each step (Figure 7) as well as biochemical affinities of TOG1 and TOG2 domains. The polarized unfurling model indicates TOG arrays form two separate states during MT polymerase activity: an αβtubulin recruitment complex and plus-end concerted polymerization complex.
These two states denoted by the TOG square and unfurled x-ray structures, which we effectively trapped by regulating the polymerization propensity for αβtubulin using DRP affinity (Figure 2 and 5). We hypothesize that the association of the αβ-tubulin bound TOG square onto the MT-plus ends, via β-tubulin binding drives the destabilization of these TOG square assemblies and promotes the concerted unfurling of the αβ-tubulins to form new protofilaments.
We envision the following steps for the polarized unfurling MT polymerase model.
First, upon binding four soluble αβ-tubulins, the dimeric TOG1-TOG2 array organizes into TOG square assemblies ( Figure 8A). These assemblies place αβtubulins in close proximity in a near head-to-tail orientation, while inhibiting their spontaneous polymerization-a feature critical for MT polymerase activity. The TOG square structure reveals that the polarized orientations of recruited αβtubulins are due to the asymmetry in each TOG domain:αβ-tubulin interface and the unique head-to-tail assembly of the dimeric TOG1-TOG2 array formed in the TOG square ( Figure 2). Positively charged unstructured SK-rich and coiled-coil dimerization regions following the TOG domains may stabilize the TOG square in the wt-Alp14 dimer (Widlund et al., 2011). The propensity of TOG1-TOG2 subunits to form head-to-tail self-assembly is likely strongly enhanced via their dimerization through their c-terminal coiled-coil. In the mammalian proteins such as XMAP215, TOG3-TOG4 likely substitutes for the second subunit of TOG1-TOG2 dimer and may play only a structural role in stabilizing the square conformation (Widlund et al., 2011). Our cysteine crosslinking/massspectrometry and mutagenesis studies indicate wt-Alp14-dimer form TOG squares readily in solution, via interfaces 1 and 2. Biochemical and negative stain-EM studies of three mutants (INT1, INT2 and INT12) suggest that interfaces 1 & 2 are responsible for TOG square organization and but they do not influence αβ-tubulin binding. Disabling these interfaces likely leads to poor preorganization of αβ-tubulins and to some degrees of spontaneous αβ-tubulin polymerization while bound to TOG1-TOG2 arrays.
The αβ-tubulin bound TOG square assemblies diffuse along MT lattice, mediated by the SK-rich regions with acidic tubulin C-termini exposed on MT surfaces ( Figure 8A) (Al-Bassam et al., 2012;Brouhard et al., 2008). Upon these TOG squares reaching the β-tubulin exposed at MT-plus end protofilament tips, αtubulin of the TOG1 or TOG2 bound αβ-tubulin may polymerize with β-tubulin exposed at the MT plus-end tip (Figure 8B-I; Figure 2B). The TOG1-αβ-tubulin polymerization is the most likely because of the high occupancy of αβ-tubulin on TOG1 due to its low exchange rate, compared to the lower occupancy of αβtubulin on TOG2 due to its rapid exchange rate. Atomic docking models ( Thus, we envision this MT plus-end driven TOG square destabilization likely triggers unfurling of a TOG square at MT plus-ends. One or more half-square TOG1-TOG2 subunit corner-like assemblies are then released. The corner-like state of αβ-tubulin bound TOG1-TOG2 subunits are likely unstable ( Figure 8B-II).
Interface 2 likely acts as a flexible hinge for TOG2 to freely rotate around TOG1 driven by Brownian diffusion, promoting its αβ-tubulin to polymerize catalytically onto the plus-end of the TOG1 bound αβ-tubulin ( Figure 8B-III). Our structural studies suggest two TOG1 and TOG2 bound αβ-tubulins polymerize in a single concerted unfurling event, as evidenced by a single elongated complex as seen in the unfurled x-ray structure ( Figure 5E, F). This effectively "unfurls" a single curved protofilament from two αβ-tubulins pre-aligned on the TOG1-TOG2 corner of a TOG square assembly. TOG array unfurling activity is directly driven by Brownian diffusion and is captured by αβ-tubulin polymerization through forming inter-dimer polymerization interfaces as shown in our structural studies ( Figure   6A, B). The αβ-tubulin inter-dimer interface (1650-Å surface area in a single interface) is fairly sufficient outcompete with TOG square (1930 Å in total for TOG square) reformation.
The unfurled TOG1-TOG2 αβ-tubulin assembly structure reveals that a gradient in αβ-tubulin affinity and exchange rate exists across TOG1 and TOG2 ( Figure   1). This gradient is positioned spatially across lower and upper positions of the newly formed protofilament with respect to the MT plus end. The tighter binding affinity of TOG1 for αβ-tubulin likely anchors the TOG array onto the MT plus end, while the rapid exchange of αβ-tubulin by TOG2 likely drives polymerization followed by release. The polarized unfurling model suggests that TOG1 and TOG2 serve specific roles in the MT polymerase activity. This is in contrast to random or reversed orientations that were suggested by other groups (Ayaz et al., 2014;Fox et al., 2014). In a newly unfurled protofilament TOG2 dissociates from upper αβ-tubulin, rapidly, while TOG1 remains tightly bound to the lower αβ-   Das et al., 2015). Taken together, our data provide a new model for a multi-step MT polymerase mechanism that accelerates αβ-tubulin polymerization.

Protein expression and purification of Alp14 and sk-Alp14 proteins
The coding regions for MT polymerases from Saccharomyces cerevisiae Stu2p Generally, constructs were transformed and expressed in BL21 bacterial strains using the T7 expression system, and were grown at 37°C and induced with 0.5 mM isopropyl thio-β-glycoside at 18 °C overnight. Cells were centrifuged and then lysed using a microfluidizer (Avastin) Extracts were clarified via centrifugation at 18,000 x g. Proteins were purified using Ni-IDA (Macherey-Nagel) and/or ion exchange using Hitrap-SP or Hitrap-Q chromatography followed by size exclusion chromatography using a Superdex 200 (30/1000) column (GE Healthcare). DRP was synthesized (Gene Art, Life Technologies), inserted into bacterial expression vectors with a C-terminal 6Xhis tag, and expressed as described above. Proteins were purified using Ni-NTA (Macherey-Nagel) followed by Hitrap Q ion exchange and followed by size exclusion chromatography as described above. Purified proteins were used immediately or frozen in liquid nitrogen for future use.
X-ray diffraction and structure determination of sk-Alp14:αβ-tubulin assemblies More than 100 sk-Alp14-monomer:αβ-tubulin: DRP crystals were screened for Xray diffraction at the Argonne National Laboratory at the Advanced Photon Source microfocus 24-ID-C beamline. Anisotropic X-ray diffraction data were collected for the best cube-shaped crystals in the P2 1 space group to 4.4 Å resolution in the best dimension, with unit cell dimensions a=216 Å, b=109 Å, and c=280 Å (Figure 2 Supplement 1). The sk-Alp14-monomer-SL:αβ-tubulin-DRP crystals showed improved diffraction and decreased anisotropy to 3.6-Å resolution in an identical P2 1 unit cell (Table S3). X-ray diffraction data were indexed and scaled using iMOSFLM and treated for anisotropic diffraction using ellipsoidal truncation on the UCLA server (services.mbi.ucla.edu/anisoscale).
Phase information was determined using TOG1 (PDB ID: wheel-like models were averaged using non-crystallographic symmetry and then refined using the PHENIX program (Adams et al., 2010). Initially, models were refined using non-crystallographic symmetry (16 fold NCS) restraints and strictly constrained coordinates with group B-factor schemes. In the final stage refinement, the strategy was switched to individual positional and isotropic Bfactor with automatic weight optimization. A 4.4-Å sk-Alp14-monomer:αβ-tubulin: DRP structure and 3.6-Å sk-Alp14-monomer-SL:αβ-tubulin:DRP structure are reported; refinement statistics appear in Table S3.
Rectangular crystals formed from sk-Alp14-monomer:αβ-tubulin: DRPΔN diffracted to 3.3-Å resolution at the Argonne National Laboratory at the Advanced Photon Source microfocus APS 24-IDC beamline. X-ray diffraction data were indexed in the P2 1 space group with unique unit cell dimensions a=115 Å, b=194 Å, and c=149 Å, with two complexes in each unit cell (table S3). Phase information was determined using molecular replacement using the TOG1, TOG2 domains and curved αβ-tubulin as search models ( Figure 5 Supplement 1B).
TOG1 and TOG2 domains were identified after cycles of density modification as described above. Four αβ-tubulins, four TOG domains, and two DRPΔN models were placed in the unit cell. The identity of TOG domains was determined using the conserved C-terminal linker and jutting helix in the TOG1 domain sequence.
A single DRPΔN molecule was identified bound per two αβ-tubulins polymerized complex. Data from each extended assembly were combined using noncrystallographic symmetry (8 fold NCS) and were averaged and refined using the program PHENIX (Adams et al., 2010) (Table S3). The individual positional coordinates and isotropic B-factor were refined with automatic weight optimization in the final stage. A 3.3-Å resolution refined density map is presented in Figure 5 Supplement 1C. Examining data quality of sk-Alp14monomer:αβ-tubulin: DRP or sk-Alp14-monomer:αβ-tubulin: DRP-ΔN using PHENIX (Adams et al., 2010) indicated the diffraction data contained a small degree of pseudo-merohedral twinning. The twin fractions were adjusted during refinement of both models.

Cysteine-crosslinking analyses of sk-Alp14 interfaces
Based on the sk-Alp14-monomer:αβ-tubulin: DRP crystal structure, S. kluyveri (sk-Alp14) ortholog protein, in its dimer form (residues 1-724), was used to generate crosslinking mutations. Interface 1 residues, which are in close proximity to each other, were mutated to cysteine: Ser180Cys (S180C) and Leu304Cys (L304C), which we termed S180C-L304C. Interface 2 residues, which are in close proximity to each other, were also mutated to cysteine: Ser41Cys (S41C) and Glu518Cys (E518C), which we termed S41C-E518C. The S. kluyveri ortholog dimer S180C-L304C mutant and S41C-E518C mutant proteins were purified as described above ( Figure 3E). These constructs were either used directly or used to make complexes with αβ-tubulin in a 2:4 (subunit:αβ-tubulin) molar ratio, as described in Figure 1A. These S180C-L304C and S41C E518C mutants, or their αβ-tubulin complexes were then treated using 5 mM Cu-phenanthroline in 50 mM HEPES and 100 mM KCl, pH 7.0 for 5 min, then treated with 5 mM EDTA. These protein mixtures were subjected to sodium dodecyl sulfate polyacrylamide gel electrophoresis under oxidizing conditions.
For LC/MS-MS mass-spectrometry based disulfide peptide mapping, S180C-L304C sk-Alp14 oxidized SDS-PAGE bands were subjected to in-gel proteolysis using either Trypsin or Chymotrypsin. Fragmented peptides were then purified and treated with 5 mM iodioacetamide, which covalently adds 57-Da in mass onto reduced cysteine containing peptides, and does not affect cysteines locked in disulfides. The peptide mixture was then treated with 5 mM Dithiothriatol to reduce disulfides and then treated with 5-Vinyl chloride, which covalently adds 105-Da mass units onto newly reduced cysteine containing peptides. LCMS/MS Mass-spectrometry was performed and the resulting peptides were analyzed.
Peptides covering 90% the sk-Alp14 were identified and majority of cysteines were identified. Only two peptides were identified with cysteine-residues that include 105-Da mass units added as described in Figure 4, S6D.

Negative stain electron microscopy of TOG-array:αβ-tubulin
SEC purified αβ-tubulin complexes of wt -Alp14-dimer, INT1, INT2 and INT12 were placed on glow discharged grids and blot after 30-60 seconds and then stained with multiple washes of 0.1% Uranyl Formate at pH 7. Grids were imaged using low-dose mode at 200 KeV using a DE-20 direct electron detector device (DDD) operating in integration mode.

Animating the MT polymerase "polarized Unfurling" mechanism
The animation was created using BLENDER 3D-animation software (http://blender.org) as follows: Briefly, surface and ribbon models of PDB coordinates representing the structures, were exported from UCSF-Chimera and were imported into BLENDER, and were then smoothed and optimized to generate cartoon models. Additional protein SK-rich regions and coiled-coil domain structures are unknown structure were thus modeled using sequence length and other information as guidance. The microtubule lattice was modeled based on the tubulin structure (PDB ID 36JF). The dissociation of TOG1 and TOG2 domains from αβ-tubulins were simulated in the animation, based on biochemical studies described in Figure 1 and S1-S3.