1. Cell Biology
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The transition state and regulation of γ-TuRC-mediated microtubule nucleation revealed by single molecule microscopy

  1. Akanksha Thawani
  2. Michael J Rale
  3. Nicolas Coudray
  4. Gira Bhabha
  5. Howard A Stone
  6. Joshua W Shaevitz
  7. Sabine Petry  Is a corresponding author
  1. Department of Chemical and Biological Engineering, Princeton University, United States
  2. Department of Molecular Biology, Princeton University, United States
  3. Department of Cell Biology, New York University School of Medicine, United States
  4. Department of Mechanical and Aerospace Engineering, Princeton University, United States
  5. Lewis-Sigler Institute for Integrative Genomics, United States
  6. Department of Physics, Princeton University, United States
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Cite this article as: eLife 2020;9:e54253 doi: 10.7554/eLife.54253

Abstract

Determining how microtubules (MTs) are nucleated is essential for understanding how the cytoskeleton assembles. While the MT nucleator, γ-tubulin ring complex (γ-TuRC) has been identified, precisely how γ-TuRC nucleates a MT remains poorly understood. Here, we developed a single molecule assay to directly visualize nucleation of a MT from purified Xenopus laevis γ-TuRC. We reveal a high γ-/αβ-tubulin affinity, which facilitates assembly of a MT from γ-TuRC. Whereas spontaneous nucleation requires assembly of 8 αβ-tubulins, nucleation from γ-TuRC occurs efficiently with a cooperativity of 4 αβ-tubulin dimers. This is distinct from pre-assembled MT seeds, where a single dimer is sufficient to initiate growth. A computational model predicts our kinetic measurements and reveals the rate-limiting transition where laterally associated αβ-tubulins drive γ-TuRC into a closed conformation. NME7, TPX2, and the putative activation domain of CDK5RAP2 do not enhance γ-TuRC-mediated nucleation, while XMAP215 drastically increases the nucleation efficiency by strengthening the longitudinal γ-/αβ-tubulin interaction.

Introduction

Microtubules (MTs) enable cell division, motility, intracellular organization and transport. Half a century ago, MTs were found to be composed of αβ-tubulin dimers, yet how MTs are nucleated in the cell to assemble the cellular structures remains poorly understood (Petry, 2016; Wu and Akhmanova, 2017). The universal nucleator, γ-tubulin efficiently nucleates MTs in vivo (Oakley and Oakley, 1989; Hannak et al., 2002; Groen et al., 2009) by forming a 2.2 megadalton, ring-shaped complex with γ-tubulin complex proteins (GCPs), known as the γ-Tubulin Ring Complex (γ-TuRC) (Moritz et al., 1995; Moritz et al., 1998; Zheng et al., 1995; Kollman et al., 2010; Kollman et al., 2015; Oegema et al., 1999). Structural studies (Kollman et al., 2010; Moritz et al., 2000; Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020) have revealed that γ-TuRC positions a lateral array of 13 γ-tubulin molecules that are thought to template MT assembly by binding αβ-tubulin dimers and promoting their lateral interaction to result in nucleation of a MT (Kollman et al., 2010; Moritz et al., 2000; Keating and Borisy, 2000; Wiese and Zheng, 2000; Kollman et al., 2011). Despite this model being widely accepted, MT nucleation from γ-TuRC molecules has not been directly visualized in real time and the dynamics of nucleation of a MT from αβ-tubulin dimers remains to be characterized. In particular, determining the critical nucleus, that is the rate-limiting transition state, for γ-TuRC nucleation is of tremendous interest, as it has important implications for how MT nucleation is spatiotemporally regulated in the cell (Figure 1A).

Figure 1 with 1 supplement see all
Single molecule microscopy of microtubule nucleation from γ-TuRC.

(A) Schematic for microtubule nucleation from γ-TuRC. Biochemical features of γ-TuRC including the γ-/αβ-tubulin interaction affinity and conformation of γ-TuRC determine to MT nucleation activity and transition state. (B) Purified, biotinylated γ-TuRC molecules were attached, incubated with 14 μM αβ-tubulin and time-lapse of MT nucleation after is shown. MTs already nucleated in the first frame are marked with yellow arrow, while new MT nucleation events between the first and last frame with blue arrows. (C) Three representative kymographs of (left) unlabeled γ-TuRC nucleating MTs colored in grayscale, or (right) fluorescently-labeled γ-TuRC, pseudo-colored in green, nucleating MTs, pseudo-colored in red. Arrows point to nucleation sites. The experiments with unlabeled γ-TuRC were repeated more than 10 times with independent γ-TuRC preparations, while those with fluorescent γ-TuRC repeated were repeated six times with three independent γ-TuRC preparations. See Figure 1—figure supplement 1 and Videos 12.

In the absence of γ-TuRC, MTs can also nucleate spontaneously from high concentrations of αβ-tubulin in vitro. In this process, which displays a nucleation barrier, the assembly of many αβ-tubulin dimers is thought to occur to form lateral and longitudinal contacts (Voter and Erickson, 1984; Flyvbjerg et al., 1996; Portran et al., 2017; Roostalu and Surrey, 2017). It has long been speculated whether γ-TuRC-mediated nucleation occurs similarly, or follows a distinct reaction pathway (Kollman et al., 2011; Roostalu and Surrey, 2017; Rice et al., 2019; Wiese and Zheng, 2006; Wieczorek et al., 2015). Moreover, the structure of native γ-TuRC shows an open conformation where adjacent γ-tubulin do not form a lateral interaction (Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020), raising further questions on how the conformational mismatch impacts γ-TuRC’s nucleation activity (Figure 1A). It has been widely proposed that γ-TuRC may transition to a closed conformation during MT assembly to match the geometry of αβ-tubulin dimers arranged laterally in the MT lattice (Kollman et al., 2015; Liu et al., 2020). This transition could further provide a mode of regulation through several putative MT-associated proteins (MAPs) that have been proposed to promote a closed conformation of γ-TuRC’s (Kollman et al., 2015; Liu et al., 2020; Kollman et al., 2011) and regulate γ-TuRC’s nucleation activity (Kollman et al., 2010; Kollman et al., 2015; Liu et al., 2020; Choi et al., 2010; Liu et al., 2014; Lynch et al., 2014). Finally, the interaction affinity between γ-tubulin and αβ-tubulin and its role on MT nucleation remain unknown (Kollman et al., 2011; Rice et al., 2019; Figure 1A).

Investigating the molecular biophysics of MT nucleation by γ-TuRC at the single-molecule level and with computational modeling have the potential to address these questions. By identifying transition states and reaction intermediates during the γ-TuRC-mediated nucleation reaction, important insights into the dynamics of MT nucleation can be revealed. Yet, technical challenges in both purifying γ-TuRC at high yield, as well as the inability to visualize MT nucleation events from individual γ-TuRC molecules in real time and at high resolution, have posed limitations. In this work, we overcome these longstanding challenges to reconstitute MT nucleation from γ-TuRC and visualize the reaction live at the resolution of single molecules. We use computational models to gain further mechanistic insights into MT nucleation and to identify the molecular composition and arrangement of the rate-limiting transition state in γ-TuRC. Finally, we examine the roles of various MAPs, particularly the co-nucleation factor XMAP215, in γ-TuRC-mediated MT nucleation and comprehensively examine how specific biomolecular features govern how MT nucleation from γ-TuRC occurs.

Results

Visualizing microtubule nucleation from γ-TuRC with single molecule microscopy

To study how γ-TuRC nucleates a MT, we purified endogenous γ-TuRC from Xenopus egg extracts and biotinylated the complexes to immobilize them on functionalized glass (Figure 1—figure supplement 1A–B). Upon perfusing fluorescent αβ-tubulin, we visualized MT nucleation live with total internal reflection fluorescence microscopy (TIRFM) (Figure 1B). Strikingly, MT nucleation events occurred specifically from γ-TuRC molecules that were either unlabeled (Figure 1B and Video 1) or fluorescently labeled during the purification (Figure 1—figure supplement 1C and Video 2). Kymographs revealed that single, attached γ-TuRC molecules assembled αβ-tubulin into a MT de novo starting from zero length within the diffraction limit of light microscopy (Figure 1C), ruling out an alternative model where MTs first spontaneously nucleate and then become stabilized via γ-TuRC. By observing fiduciary marks on the MT lattice (Figure 1C) and generating polarity-marked MTs from attached γ-TuRC (Figure 1—figure supplement 1D), we show that γ-TuRC caps the MT minus-end while only the plus-end polymerizes, as supported by previous works (Keating and Borisy, 2000; Wiese and Zheng, 2000). Notably, the detachment of γ-TuRC molecules and re-growth of the MT minus-ends were not observed, and γ-TuRC persists on the MT minus-end for the duration of our experiments. Altogether, our results demonstrate that γ-TuRC directly nucleates a MT.

Video 1
Microtubule nucleation from γ-TuRC complexes.

γ-TuRC was attached to functionalized coverslips and MT nucleation was observed upon introducing fluorescent αβ-tubulin (gray). MTs nucleated from individual γ-TuRC molecules from zero length at 14 μM αβ-tubulin and the plus-end of nucleated MTs polymerized, but not its minus-end. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

Video 2
Microtubule nucleation from fluorescent, single γ-TuRC molecules.

Dual Alexa-568 and biotin-labeled γ-TuRC (green) was attached to functionalized coverslips and MT nucleation was observed upon introducing fluorescent αβ-tubulin (red). MTs nucleated from single γ-TuRC molecules at 10.5 μM αβ-tubulin. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

Molecular composition of the transition state during γ-TuRC-mediated nucleation

To determine how γ-TuRC nucleates a MT, we measured the kinetics of MT nucleation for a constant density of γ-TuRC molecules and increasing αβ-tubulin concentrations (Figure 2A and Video 3). γ-TuRCs nucleated MTs starting from 7 μM tubulin (Figure 2A–B), which is higher than the minimum tubulin concentration (C*) needed for growth from a pre-formed MT plus-end (C*=1.4 μM, Figure 2B). Furthermore, the number of MTs nucleated from γ-TuRC increased non-linearly with αβ-tubulin concentration as opposed to the linear increase in MT’s growth speed with tubulin concentration (Figure 2B). By measuring the number of MTs nucleated over time with varying αβ-tubulin concentration (Figure 2C), we calculated the rate of MT nucleation. The power-law dependence on αβ-tubulin concentration (Figure 2D) yields the number of tubulin dimers, 3.9 ± 0.5, that compose the rate-limiting, transition state during MT assembly from γ-TuRC (Figure 2D). Thus, the cooperative assembly of nearly 4 αβ-tubulin subunits on γ-TuRC represents the most critical, rate-limiting step in MT nucleation.

Molecular composition of transition state in γ-TuRC-mediated nucleation.

(A) Titrating tubulin concentration with constant the density of γ-TuRC. MT nucleation from γ-TuRC begins at 7μM tubulin. (B) MT plus-end growth speed increases linearly with tubulin concentration. Individual data points are plotted, and linear fit (red line) with shaded mean±2std (95% confidence interval) is displayed. Critical concentration for polymerization as C* = 1.4 μM. Inset: Number of MTs nucleated by γ-TuRCs within 120 s varies non-linearly with tubulin concentration. (C) Number of MTs nucleated (N(t)) over time (t) is plotted for varying tubulin concentration to obtain rate of nucleation as the slope of the initial part of the curves. Shaded regions represent 95% confidence interval (n±2n) in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. (D) Number of tubulin dimers (n) in the critical nucleus on γ-TuRC was obtained as 3.9±0.5 from the equation dNdt|t0=kCtubn displayed on a log-log axis as detailed in Materials and methods. The rate of nucleation at 10.5 μM was set to 1 to normalize differences in γ-TuRC concentration from individual experiments. The experiments and analyses in (A–D) were repeated identically three times with independent γ-TuRC preparations. MT nucleation data, prior to normalization, from one representative dataset is displayed in (B–C). Analyses from all repeats was pooled and normalized as described above, and data points from 15 nucleation-time curves are plotted in (D). See Video 3.

Video 3
γ-TuRC molecules nucleate microtubules efficiently.

Constant density of γ-TuRC was attached while concentration of fluorescent αβ-tubulin was titrated (3.5–21 μM) and MT nucleation was observed. γ-TuRC molecules nucleated MTs starting from 7 μM tubulin and MT nucleation increased non-linearly with increasing tubulin concentration. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

γ-TuRC-mediated nucleation is more efficient than spontaneous nucleation

Based on the traditional assay where MTs are nucleated, fixed and visualized, a large variability in γ-TuRC’s MT nucleation activity has been observed. With this setup, γ-TuRC has often been reported to be a poor nucleator with a similar activity as spontaneous MT nucleation (Moritz et al., 1995; Zheng et al., 1995; Kollman et al., 2010; Kollman et al., 2015; Oegema et al., 1999; Liu et al., 2020; Wieczorek et al., 2020; Choi et al., 2010; Thawani et al., 2018). With our live TIRFM assay, we aimed to quantitatively compare the efficiency of γ-TuRC-mediated MT nucleation with spontaneous MT nucleation (Figure 3A). In contrast to γ-TuRC-mediated nucleation, a higher concentration of 14 μM tubulin was required for any spontaneous assembly of MTs, after which both the plus- and minus-ends polymerize (Figure 3B, Figure 3—figure supplement 1A and Video 4). The number of MTs assembled as a function of the αβ-tubulin concentration displayed a power-law dependence with an even larger exponent of 8.1 ± 0.9 (Figure 3C), indicating a highly cooperative process that requires 8 αβ-tubulin dimers in a rate-limiting intermediate, in agreement with previous reports (Voter and Erickson, 1984; Flyvbjerg et al., 1996). Further, direct comparison and measurement of spontaneous MT assembly with γ-TuRC-mediated nucleation (Figure 3—figure supplement 1B–C) clearly demonstrates that γ-TuRC nucleates MTs significantly more efficiently. Notably, specific attachment of γ-TuRC to coverslips is also required to observe the nucleation activity (Figure 3—figure supplement 1C). In sum, γ-TuRC-mediated nucleation occurs efficiently and its critical nucleus requires less than half the number of αβ-tubulin dimers compared to spontaneous assembly.

Figure 3 with 1 supplement see all
Comparison of γ-TuRC-mediated, spontaneous and seed-templated nucleation.

(A) Spontaneous MT nucleation (schematized) was measured with increasing tubulin concentration and high concentrations. 14 μM tubulin is required. (B) Number of MTs (N(t=τ)) nucleated spontaneously were plotted against tubulin concentration. Power-law curve was fit as N(t=τ)=kCn on a log-log axis, and linear scale in the inset. Tubulin cooperativity (exponent) of n = 8.1±0.9 was obtained as detailed in Methods. Experiments and analyses in (A–B) were repeated thrice independently, all data were pooled and data points from 11 nucleation curves are plotted in (B). In the inset, data is represented in linear plot, where shaded regions represent 95% confidence interval (n±2n) in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. Scale bars, 10μm. (C) Schematic and an example micrograph of blunt, stabilized MT seeds is shown and MT assembly from them was observed (bottom) with varying tubulin concentration. (D) Cumulative probability of MT assembly from seeds (p(t)) over time (t) is plotted and rate of nucleation was obtained as the slope from initial part of the curves. Shaded regions represent 95% confidence interval (n±2n) in the number of MTs assembled (n) from seeds as described in Materials and methods. (E) As described in Methods, the measurements fit well to equation dpdt|t0=k(CC)n displayed on a log-log plot. n = 1±0.3 was obtained showing nearly non-cooperative assembly of tubulin dimers. The experiments and analyses in (C–E) were repeated three times independently. MT nucleation data, prior to normalization, from one representative dataset is displayed in (C–D). Analyses from all experiments was pooled, and data points from a total of 11 nucleation-time curves are reported in (E). See Figure 3—figure supplement 1 and Videos 4 and 5.

Video 4
Spontaneous microtubule nucleation occurs at high tubulin concentration.

Concentration of fluorescent αβ-tubulin was titrated (7–21 μM) and spontaneous MT nucleation was assayed. MTs nucleated spontaneously starting from high concentration of 14 μM tubulin and MT nucleation increased non-linearly with tubulin concentration. Both plus- and minus-ends of the assembled MTs polymerize. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

Contribution of end architecture of γ-TuRC to microtubule nucleation

The MT plus-end architecture, which ranges from blunt to tapered, is critical for MT polymerization dynamics (Gardner et al., 2014; Mickolajczyk et al., 2019; Brouhard and Rice, 2018), and was recently proposed to be critical for MT nucleation (Wieczorek et al., 2015). To investigate how the blunt-end geometry of γ-TuRC contributes to its nucleation kinetics and transition state, we generated Alexa-568 labeled, stable MT seeds with blunt ends as described previously (Wieczorek et al., 2015) and compared MT assembly from seeds upon addition of Cy5-labelled αβ-tubulin dimers (Figure 3C) side-by-side with γ-TuRC-mediated nucleation. At a minimum concentration of 2.45 μM, approaching the critical concentration needed for polymerization of a MT plus-end, a large proportion of pre-formed MT seeds assemble MTs (Figure 3C–D, Figure 3—figure supplement 1D and Video 5). At 7 μM tubulin, the rate of assembly of MTs from the blunt seeds increased to reach the maximum rate that could be temporally resolved, that is all of the MT seeds immediately assembled a MT (Figure 4D). This is in contrast to the kinetics of γ-TuRC-mediated nucleation at 7 μM tubulin concentration, where minimal nucleation activity was observed (Figure 2C–D). The measured reaction kinetics as a function of the αβ-tubulin concentration (Figure 3D) was used to obtain the power-law of the nucleation rate, 1 ± 0.3 (Figure 3E). This suggests that in our assay condition, blunt MT ends assemble tubulin dimers into the MT lattice non-cooperatively. In other words, the addition of a single αβ-tubulin dimer suffices to overcome the rate-limiting barrier, which also occurs during the polymerization phase of MT dynamics. Notably, when this experiment was replicated with the coverslip preparation and assay conditions reported previously (Wieczorek et al., 2015), a high concentration of tubulin was necessary for seeds to assemble MTs in agreement with the previous work (Wieczorek et al., 2015). However, our assay conditions, that were used to compare seed-templated MT assembly with γ-TuRC-mediated nucleation side-by-side, result in a low, minimal tubulin concentration that is needed for seed-mediated MT assembly. To conclude, while the γ-TuRC positions a blunt plus-end of γ-tubulins, the contribution of this specific end architecture in defining the kinetics of nucleation from γ-TuRC and its transition state is minimal.

Figure 4 with 2 supplements see all
γ-tubulin binds to αβ-tubulin with a high affinity.

(A) Size-exclusion chromatography was performed with 150 nM of γ-tubulin alone (i) and with 35 μM and 10 μM αβ-tubulin in (ii) and (iii), respectively. Gel filtration fractions were analyzed via SDS–PAGE followed by immunoblot with γ-tubulin and αβ-tubulin antibodies. A shift in the γ-tubulin elution to fraction H was observed with both 35 μM and 10 μM αβ-tubulin, denoting complex formation with αβ-tubulin. See Figure 4—figure supplement 1A. Stokes’ radii of reference proteins: thyroglobulin (8.6 nm), aldolase (4.6 nm) and ovalbumin (2.8 nm), are marked at their elution peak. Size exclusion runs were repeated three times, with the exception of 10 μM αβ-tubulin run that was performed twice. (B) Single molecule microscopy was performed with γ-tubulin and αβ-tubulin. Control buffer (left panels, (i) and (ii)) or biotinylated αβ-tubulin (right panels, (i) and (ii)) was attached to coverslips, incubated with fluorescent αβ-tubulin (i) or γ-tubulin (ii) molecules, set as 0 s, and their binding at 60–90 s. (C) Number of bound molecules were analyzed for the first 15 s of observation described in Materials and methods. Experiments and analyses in (B–C) were repeated identically two times, pooled and reported. n = 56 data points each were displayed as mean ± std in the bar graph in (C). Further confirmed with a third supporting experimental set where the observation began later at 180 s and was therefore, not pooled. See also Figure 4—figure supplements 12.

Video 5
Microtubule assembly from blunt plus-ends resembles polymerization.

MTs with blunt ends (seeds, cyan) were generated and attached to functionalized coverslips. Varying concentration of fluorescent αβ-tubulin was added (1.4–8.7 μM, pseudo-colored as magenta) and MT assembly from seeds was assayed. MTs assembled at concentration above 1.4 μM tubulin, which is the minimum concentration needed for polymerization of MT plus-ends (C*). Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

In summary, because γ-TuRC positions an array of γ-tubulins at its nucleation interface that are thought to stabilize intrinsically weak, lateral αβ/αβ-tubulin interaction (Kollman et al., 2010; Kollman et al., 2015; Liu et al., 2020; Wieczorek et al., 2020; Kollman et al., 2011; Roostalu and Surrey, 2017; Rice et al., 2019), MT nucleation by γ-TuRC has been proposed to function similar to polymerization of a MT end. Here, we show several lines of evidence that γ-TuRC-mediated nucleation has distinct characteristics from MT polymerization and assembly from blunt MT seeds. While growth speed of MTs nucleated from γ-TuRC or templated from MT seeds is similar (Figure 3—figure supplement 1D), γ-TuRC molecules do not nucleate MTs at low tubulin concentration where MT polymerization can occur. Further increasing tubulin concentration results in a non-linear increase in the number of γ-TuRCs molecules that nucleate MT, as opposed to a linear increase in rate of assembly from seeds. At the highest tubulin concentrations tested, approximately 10–15% of γ-TuRCs nucleate MTs in the TIRF assays (see Materials and Methods). While these results were obtained with endogenous γ-TuRCs purified from cytosol, it remains possible that specific factors at MTOCs can modulate γ-TuRC’s conformation and kinetics. In summary, the rate-limiting transition state on γ-TuRC is composed of four αβ-tubulin dimers in contrast with MT polymerization where one tubulin dimer suffices to overcome the slowest step.

γ-tubulin has a high affinity for αβ-tubulin

Consequently, specific biochemical features of γ-TuRC must govern its nucleation activity and the composition of the transition state during nucleation. To address this, we first measured the interaction affinity between γ-tubulin and αβ-tubulin, which could provide insight into γ-TuRC’s nucleation interface and its role in MT nucleation. To begin, we performed size-exclusion chromatography where γ-tubulin alone elutes as a broad peak in fractions I-N (Figure 4A (i), pseudo-colored profile in green) at low concentration. Interestingly, in the presence of either 10 μM (low) or 35 μM (high) concentrations of αβ-tubulin, the γ-tubulin binds to αβ-tubulin (pseudo-colored profile in cyan) and elutes earlier, specifically in fraction H (Figure 4A (ii-iii), yellow arrow). Further, the overall elution profile of γ-tubulin is altered to follow αβ-tubulin, showing that γ-tubulin binds to αβ-tubulin at both the low and high concentrations we tested. To compare this with αβ-/αβ-tubulin’s longitudinal interaction, we performed chromatography of αβ-tubulin alone (Figure 4—figure supplement 1A). At a lower concentration (10 μM), αβ-tubulin elutes only as a single subunit in fractions H-K (Figure 4—figure supplement 1A (i)). Only at high αβ-tubulin concentration (35 μM) did we detect a small population of αβ-tubulin bound to another αβ-tubulin (Figure 4—figure supplement 1A (i), fractions B-C denoted with red arrows). This suggests that the heterogeneous γ-/αβ-tubulin affinity is higher than the αβ-/αβ-tubulin affinity.

To further investigate how γ-tubulin and αβ-tubulin interact, we turned to single molecule microscopy. We attached biotinylated αβ-tubulin dimers to a coverslip, added either fluorescently labeled αβ-tubulin (Figure 4B (i)) or γ-tubulin (Figure 4B (ii)) to the solution, and visualized the binding of single fluorescent molecules to αβ-tubulin molecules on the coverslip. While both fluorescent αβ-tubulin and γ-tubulin specifically bind to surface-attached αβ-tubulin, 15-fold more γ-tubulin molecules were bound than αβ-tubulin molecules (Figure 4C), further supporting a stronger γ-/αβ-tubulin interaction. Finally, these results were confirmed with a biolayer interferometry assay, where lower concentrations of γ-tubulin were detected to interact with probe-bound αβ-tubulin, while a much higher concentration of αβ-tubulin was necessary to measure an interaction between αβ-/αβ-tubulin dimers (Figure 4—figure supplement 1B). These results are congruent with in vivo γ-/αβ-tubulin affinity measurements made in yeast cells (Erlemann et al., 2012).

In performing the above experiments, we unexpectedly found that purified γ-tubulin on its own, at high concentrations and at 33°C, efficiently nucleated MTs from αβ-tubulin subunits (Figure 4—figure supplement 2A) and capped MT minus-ends while allowing plus-ends to polymerize (Figure 4—figure supplement 2B). Besides its ability to form higher order oligomers in a physiological buffer (Thawani et al., 2018), γ-tubulin at high concentrations also forms filaments in vitro of variable widths (Figure 4—figure supplement 2C; Moritz and Agard, unpublished results) as assayed by negative stain electron microscopy (EM). The formation of filaments in vitro is consistent with the previous in vivo observations where γ-tubulin was over-expressed and immunoprecipitated (Lindström and Alvarado-Kristensson, 2018; Chumová et al., 2018; Pouchucq et al., 2018). To understand the nature of these filaments, we generated 3D reconstructions, which revealed that γ-tubulins self-assemble into lateral arrays with a repeating unit of approximately 54 Å (Figure 4—figure supplement 2D–E). This closely matches the lateral tubulin repeats in the MT lattice (PDB:6DPU [Zhang et al., 2018; Zhang and Nogales, 2018]) and in γ-tubulin crystal contacts (52 Å, PDB:1Z5W [Aldaz et al., 2005a; Aldaz et al., 2005b]), but not the longitudinal αβ-tubulin repeat (40 Å). This suggests that laterally associated γ-tubulin are sufficient to efficiently nucleate MTs.

In sum, at the nucleation interface of γ-TuRC, γ-tubulin has a higher longitudinal affinity for αβ-tubulin compared to αβ-tubulin’s affinity for itself, which promotes MT nucleation from γ-TuRC.

Monte Carlo simulations recapitulate the dynamics microtubule nucleation from γ-TuRC

To further probe the dynamics of MT nucleation, we developed Monte Carlo simulations to model MT nucleation from γ-TuRC. Our model was based on one previously developed for the plus-end dynamics of a MT (Mickolajczyk et al., 2019; VanBuren et al., 2002; Ayaz et al., 2014). A 13-protofilament geometry for the MT lattice and γ-TuRC were used with a pitch of 3 tubulins (Figure 5A). αβ-tubulin dimers arrive with a constant on rate, kon (μM−1s−1) on each protofilament. The interactions between αβ-tubulins was assumed to occur with longitudinal and lateral bond energies, GLong,αβ-αβ and GLat,αβ-αβ, respectively, similar to previous literature (Mickolajczyk et al., 2019; VanBuren et al., 2002; Ayaz et al., 2014). The longitudinal bond energy between γ-/αβ-tubulin, GLong,γ-αβ determines the dwell time of αβ-tubulin dimers on γ-TuRC. An open conformation of native γ-TuRC was assumed, as observed in recent structural work (Liu et al., 2020; Wieczorek et al., 2020), where lateral interactions between tubulins on neighboring sites were not allowed. A thermodynamic barrier,  GγTuRC-conf and a pre-factor rate constant kγTuRC-conf (s−1) determine the transition from this open to closed γ-TuRC conformation where lateral tubulin interactions can occur (Figure 5A). As αβ-tubulin dimers assemble on γ-TuRC, the free energy of this transition decreases by the total energy of all n lateral bonds that can be formed, GγTuRC-conf-nGLat,αβ-αβ.

Figure 5 with 2 supplements see all
Monte Carlo simulations of microtubule nucleation from γ-TuRC.

(A) Kinetic Monte Carlo simulations of MT nucleation were performed. Helical MT lattice was simulated with 13 protofilaments and a pitch of 3 tubulin monomers across the seam. Native γ-TuRC was simulated in an open conformation and was allowed transition into a closed conformation with a thermodynamic penalty of GγTuRC-conf. αβ-tubulin dimers form longitudinal bonds with energies, GLong,γ-αβ and GLong,αβ-αβ to γ-tubulin and other αβ-tubulins, respectively, and lateral bond with energy, GLat,αβ-αβ with neighboring αβ-tubulin dimers. (B) MT length (μm) versus time (seconds) traces of two independent simulations are presented (bottom). MT nucleation occurs are variable time points for each model realization. Zoomed-in insets of the first simulation show the length of the tallest protofilament (nm) and total number of αβ-tubulin dimers assembled in the first 200 ms and 5 s near the transition state of the simulation. (C) Simulations were performed with kon=1.3×106 (M1s1pf1)ΔGLong,αβαβ=7.2kBT, ΔGLat,αβαβ=6.5kBT, ΔGLong,γαβ=1.1ΔGLong,αβαβ, kγTuRC-conf=0.01s-1. ΔGγTuRCconf was varied from +(0-30)kBT. Tubulin concentration was varied from 2.5 to 50 μM. 200 simulations were performed for a given tubulin concentration at every parameter set, except for ΔGγTuRCconf=10kBT where 500 simulations were performed. From probability of MT nucleation (p(t)) versus time (t) curves, the initial rate of nucleation dpdt|t0 was measured and plotted against concentration on a log-log axis as detailed in Materials and methods. (D) With the parameters defined above and ΔGγTuRCconf=10kBT, the transition state at the time of γ-TuRC’s conformational change was recorded for n=2119 simulations. Normalized histogram of total number of αβ-tubulin dimers is plotted (left). Three-dimensional probability distribution of total number of αβ-tubulin dimers (x) and number of lateral αβ-tubulin interactions (y) is plotted (right). The most populated transition states are denoted with coordinates (x,y) and schematized. See also Figure 5—figure supplements 1, 2.

MT growth parameters were determined by fitting to experimental growth speed curves (Figure 5—figure supplement 1A) and were found to be similar to previous estimates (Mickolajczyk et al., 2019; VanBuren et al., 2002). Based on our biochemical measurement (Figure 5B), GLong,γ-αβ was estimated to be higher than GLong,αβ-αβ, while a wide range was explored for the other parameters. The resulting model produces a sharp transition from zero-MT length to a continuously growing MT upon γ-TuRC closure (Figure 5B) that occurs at variable time points for each realization of the model (Figure 5B and Figure 5—figure supplement 1B-C(i)). This qualitatively recapitulates the dynamics of γ-TuRC-mediated nucleation events observed experimentally.

Nucleation kinetics and the power-law dependence on αβ-tubulin concentration was obtained by simulating hundreds of model realizations. While kγTuRC-conf and ΔGLong,γαβ do not alter the power-law exponent significantly, they set the rate of nucleation at a specific αβ-tubulin concentration (Figure 5—figure supplement 1B-C). The thermodynamic barrier, ΔGγTuRCconf instead determines the power-law exponent and the number of αβ-tubulins in the rate-limiting, transition state (Figure 5C). At ΔGγTuRCconf<2.5kBT, cooperative assembly of 1-2 αβ-tubulins suffice to nucleate MTs, while at high ΔGγTuRCconf>20kBT, more than 5 αβ-tubulins assemble cooperatively for successful MT nucleation. At an intermediate ΔGγTuRCconf=10kBT, MT nucleation kinetics and its power-law dependence recapitulates our experimental measurements (compare Figure 5—figure supplement 2A with Figure 2C-D). Here, γ-TuRCs minimally nucleate MTs at 7μM tubulin, MT nucleation increases non-linearly with tubulin concentration, and 4 ± 0.4 αβ-tubulins compose the transition state (Figure 5—figure supplement 2A and Figure 5C, green curve highlighted with an asterisk).

As a further validation of our model, we simulated the dynamics of MT nucleation from blunt MT seeds. Here, we assumed that MT assembly begins from a closed γ-TuRC geometry where all longitudinal bond energies were set equal to GLong,αβ-αβ (Figure 5—figure supplement 2B). The simulations predict near complete MT assembly at minimal αβ-tubulin concentration of 2μM and transition state of 1.1 ± 0.1 αβ-tubulins (Figure 5—figure supplement 2A), in agreement with MT assembly from blunt seeds that we measured experimentally (Figure 3D-E). Thus, our Monte Carlo simulations accurately capture the detailed dynamics of MT nucleation from γ-TuRC.

Arrangement of αβ-tubulin dimers in transition state for γ-TuRC-mediated microtubule nucleation

We next characterized the dynamics of αβ-tubulins during MT nucleation from γ-TuRC by examining the time traces from individual model simulations. First, prior to MT nucleation, we observe longitudinal association of individual αβ-tubulins either to the γ-tubulin sites on the open γ-TuRC or, less frequently, with existing αβ-tubulin in a protofilament (Figure 5B, left insets). These αβ-tubulins dissociate rapidly in the absence of additional lateral bond energy. Once a MT lattice is assembled, persistence of αβ-tubulin dimers with both longitudinal and lateral contacts drive the growth of plus-end. Analogous observations during growth of a MT plus-end also show rapid dissociation of αβ-tubulin that form only a longitudinal contact, while ones with additional lateral contacts persist (Mickolajczyk et al., 2019). At the sharp transition prior to MT assembly (Figure 5B, right insets), we find that many αβ-tubulin dimers stochastically assemble on neighboring sites on γ-TuRC. Favorable Gibbs free energy from the lateral interaction between these αβ-tubulin dimers overcomes the energy penalty of the conformational change and transitions γ-TuRC into a closed state.

Finally, we characterize the arrangement of αβ-tubulin dimers in the rate-limiting, transition state that results in a closed γ-TuRC conformation prior to MT polymerization. A variable total number of αβ-tubulin dimers with an average of 5.2 ± 1 (n = 2119 simulations) were present on γ-TuRC at the transition state (Figure 5D, left). To our surprise, αβ-tubulin subunits in the transition state assemble on neighboring sites into laterally arranged groupings (Figure 5D, right). The most probable transition state is composed of four αβ-tubulin arranged on neighboring sites that form three lateral bonds when the γ-TuRC conformation changes to a closed one. The other probable states have 5 αβ-tubulins arranged laterally in two groups of 2 and 3 dimers each, or in two groups of 1 and 4 dimers each, and 6 αβ-tubulins arranged in two groups of 2 and 4 dimers, or in two groups of 3 dimers each. Most importantly, in these transition states, the free energy gained from the lateral bonds between αβ-tubulins compensates for the thermodynamic barrier posed by γ-TuRC’s open conformation to allow for MT nucleation. Notably, the laterally-arranged group of 4 αβ-tubulin dimers physically represents the power-law exponent measured from the average nucleation kinetics (Figure 2D).

Role of putative activation factors in γ-TuRC-mediated nucleation

Next, we investigated how accessory factors regulate γ-TuRC-dependent MT nucleation. While several activation factors (Choi et al., 2010; Liu et al., 2014; Alfaro-Aco et al., 2017) have been proposed to enhance the MT nucleation activity of γ-TuRC, the function of these putative activation factors remains to be tested with a sensitive and direct assay. We incubated the purified γ-TuRC activation domain (γ-TuNA) (Choi et al., 2010) from Xenopus laevis protein CDK5RAP2 with γ-TuRC at high concentrations to maximally saturate the binding sites on γ-TuRC (Figure 6A), and further supplemented additional γ-TuNA with αβ-tubulin used during the nucleation assay. Measurement of nucleation activity revealed that CDK5RAP2’s γ-TuNA domain increases γ-TuRC-mediated nucleation only by 1.4 (±0.02) -fold (mean ± std, n = 2) at t = 180 s, falling within the 95% confidence intervals of the control reactions (Figure 6A–B and Video 6). Another putative activator, NME7 (Liu et al., 2014), when added to γ-TuRC at saturating concentrations (Wühr et al., 2014; Figure 6—figure supplement 1 and Video 6), did not increase γ-TuRC’s nucleation activity (Figure 6B). Finally, we assessed the protein TPX2 that not only contains a split γ-TuNA and overlapping SPM (Alfaro-Aco et al., 2017), but also functions as an anti-catastrophe factor in vitro (Wieczorek et al., 2015; Roostalu et al., 2015) and was proposed to stimulate γ-TuRC-mediated nucleation (Alfaro-Aco et al., 2017; Tovey and Conduit, 2018; Zhang et al., 2017). TPX2 also had a small increase on the nucleation activity of γ-TuRC by 1.2 (±0.3) -fold (mean ± std, n = 3) at t = 180 s, but bound strongly along the MT lattice (Figure 6C–D and Video 6). While high concentration of TPX2 forms condensates with αβ-tubulin and promotes spontaneous MT nucleation (Roostalu et al., 2015; King and Petry, 2020), near its endogenous concentration of TPX2 (Thawani et al., 2019) used here, TPX2 is able to saturates the MT lattice, yet it does not significantly increase γ-TuRC-mediated nucleation, in agreement with the physiological observations (Alfaro-Aco et al., 2017). Thus, the putative activation motif of CDK5RAP2, full-length NME7 or TPX2 all have minor effects on γ-TuRC’s MT nucleation activity.

Figure 6 with 1 supplement see all
Regulation of γ-TuRC-mediated nucleation by putative activation factors.

(A) A constant density of γ-TuRC molecules were attached without (left) and with (right) 6μM CDK5RAP2’s γ-TuNA motif and 10.5μM tubulin ± 3μM additional γ-TuNA was added. Scale bar, 10 μm. (B) MTs nucleated from γ-TuRC molecules were analyzed and 3-6 μM CDK5RAP2’s γ-TuNA motif (left) or 1-6 μM NME7 (right). Experiments and analyses in (A–B) were individually repeated twice on different days of experimentation with independent or same γ-TuRC preparations. Number of MTs nucleated in control reactions at 200 s for γ-TuNA, and at 150 s for NME7 was set to 1 to account for variable γ-TuRC concentration across purifications, all data were pooled and reported. Individual datasets with ±γ-TuNA and ±NME7 is represented with solid or dashed curves. Shaded regions represent 95% confidence interval (n±2n) from each dataset in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. (C–D) A constant density of γ-TuRC molecules were attached and 10.5 μM tubulin ± 10-20 nM GFP-TPX2 was added. Experiments and analyses were repeated thrice with independent γ-TuRC preparations. To account for the variable γ-TuRC concentration across purifications, the number of MTs nucleated in control reactions at 150 s was set to 1. All data were pooled and reported. Individual dataset with ±TPX2 is represented with solid, dashed or dotted curves. Shaded regions represent 95% confidence interval (n±2n) from each dataset in the number of nucleated MTs (n) assuming a Poisson distribution as described in Methods. See Fiure 6—figure supplement 1 and Video 6.

Video 6
γ-TuNA motif from CDK5RAP2, NME7 and TPX2 do not significantly increase γ-TuRC-mediated microtubule nucleation.

Top panels: γ-TuRC was immobilized on coverslips with control buffer (left) or with 6 μM γ-TuNA motif from CDK5RAP2 (right) and MT nucleation was observed upon introducing fluorescent 10.5 μM αβ-tubulin (gray) without or with 3 μM γ-TuNA, respectively. Middle panels: γ-TuRC was immobilized on coverslips with control buffer (left) or with 6 μM NME7 (right) and MT nucleation was observed upon introducing fluorescent 10.5 μM αβ-tubulin (gray) without or with 1 μM NME7, respectively. Bottom panels: γ-TuRC was immobilized on coverslips and MT nucleation was observed upon introducing fluorescent 10.5 μM αβ-tubulin (pseudo-colored as red) without or with 10 nM GFP-TPX2 (right, labeled as green). TPX2 bound along the nucleated MTs but did not significantly increase the MT nucleation activity of γ-TuRC molecules. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

XMAP215 promotes microtubule nucleation by strengthening the longitudinal bond energy between γ-TuRC and αβ-tubulin

Recently, XMAP215 was discovered to be a nucleation factor that synergizes with γ-TuRC in X. laevis and S. cerevisiae (Thawani et al., 2018; Gunzelmann et al., 2018), or works in an additive manner with γ-tubulin (King et al., 2020). To investigate how XMAP215 participates in MT nucleation, we performed single molecule experiments with XMAP215 and γ-TuRC. At low tubulin concentrations of 3.5 μM and 7 μM, where none or little MT nucleation occurs from γ-TuRCs alone (Figure 7A and Figure 7—figure supplement 1A), as shown earlier. Strikingly, the addition of XMAP215 induced many surface-attached γ-TuRCs to nucleate MTs, resulting in a drastic increase in number of nucleated MTs by 25 (±9) -fold (mean ±std, n = 3) within t = 120 s (Figure 7A–B, Fig. Figure 7—figure supplement 1B and Video 7). By directly visualizing γ-TuRC and XMAP215 molecules during the nucleation reaction (Figure 7C), we found that XMAP215 and γ-TuRC molecules first form a complex from which a MT was then nucleated (Figure 7C and Video 8). For 76% of the events (n = 56), XMAP215 visibly persisted between three to ≥300 s on γ-TuRC before MT nucleation. After MT nucleation, XMAP215 molecules polymerize and track with the MT plus-end. For 50% of nucleation events (n = 58), some XMAP215 molecules remained on the minus-end together with γ-TuRC, while for the other 50% of events, XMAP215 was not observed on the minus-end after nucleation. This suggests that XMAP215 molecules nucleate with γ-TuRC and then continue polymerization of the plus-end.

Figure 7 with 2 supplements see all
Role of XMAP215 and microtubule-associated proteins in microtubule nucleation with γ-TuRC.

(A) γ-TuRCs were attached and 7μM tubulin (pseudo-colored in red) ± 20nM XMAP215-GFP (pseudo-colored in green) was added. Scale bar, 10 μm. Experiments and analyses in (A–B) were repeated thrice with independent γ-TuRC preparations. (B) Number of MTs nucleated (N(t)) over time (t) was measured and control reactions at 120 s was set to 1 to account for variable γ-TuRC concentration across purifications, all data were pooled and reported. Individual datasets with ±XMAP215 is represented with solid or dashed curves. Shaded regions represent 95% confidence interval (n±2n) from each dataset in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. See also Figure 7—figure supplement 1A-B. (C) Sequence of events during cooperative MT nucleation by γ-TuRC and XMAP215 was visualized using labeled γ-TuRC (blue), XMAP215 (red) and tubulin (green) represented in a time sequence and kymograph. γ-TuRC and XMAP215 form a complex prior to MT nucleation. XMAP215 molecules reside on γ-TuRC for before MT nucleation. The experiment was repeated a total of eight times with two independent γ-TuRC preparations and independent XMAP215 purifications. Scale bar, 5μm. (D–E) Number of MTs nucleated (N(t)) over time (t) was measured after titrating tubulin with constant γ-TuRC and XMAP215 concentration. XMAP215/γ-TuRC molecules nucleate MTs from 1.6 μM tubulin. Shaded regions represent 95% confidence interval (n±2n) in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. (E) Number of tubulin dimers (n) in the critical nucleus on cooperative nucleation by γ-TuRC/XMAP215 was obtained as 3.3 ± 0.8 from the equation dNdt|t0=kCtubn displayed on a log-log axis as detailed in Materials and methods. The rate of nucleation at 3.5μM was set to 1 to normalize differences in γ-TuRC concentration from individual experiments. Experiment and analyses in (D–E) was repeated thrice over the entire concentration range with independent γ-TuRC preparations, and fewer concentration points were repeated another two times. All five datasets were pooled and data points from a total of 18 nucleation-time curves are reported in (E). Simulations were adapted to understand how XMAP215 changes the thermodynamics of γ-TuRC-mediated nucleation. Parameter values used: kon=1.3×106 M-1s-1pf-1, GLong,αβ-αβ=-8.64kBT, GLat,αβ-αβ=-6.2kBT, GLong,γ-αβ=-9.5kBT, kγTuRC-conf=0.01s-1 and  GγTuRC-conf=10kBT. Compared to simulations for γ-TuRC alone (Figure S6A), either GLong,αβ-αβ was increased 1.2-fold, as proposed previously (VanBuren et al., 2002), or both GLong,αβ-αβ and GLong,αβ-αβ were increased 1.2-fold. 200 simulations each were performed for a range of tubulin concentration 1.6-7 μM. Probability of MT nucleation (p(t)) versus time (t) is displayed in (D). The initial rate of nucleation dpdt|t0 was measured at each tubulin concentration and plotted against concentration on a log-log axis in (E). Linear curve was fit for n=5 simulated data points, and critical nucleus of 3.8 ± 0.3 αβ-tubulins. Increasing all longitudinal bond energies reproduces the effect of XMAP215 on γ-TuRC-mediated nucleation. (F) Number of MTs nucleated was measured to assess the effect of inhibitory MAPs MCAK or Stathmin on γ-TuRC-mediated nucleation. 10.5μM tubulin ± 10nM MCAK, or 7-10.5μM tubulin ± 2-5μM Stathmin was added to attached γ-TuRC- molecules, and MCAK and Stathmin were both found to inhibit γ-TuRC-mediated nucleation. Experiments and analyses for both MAPs were repeated thrice individually with independent γ-TuRC preparations. Number of MTs nucleated in control reactions at 200 seconds was set to 1 to account for variable γ-TuRC concentration across purifications, all data was pooled and reported. Individual dataset with ± MCAK are reported with solid, dashed or dotted curves. For Stathmin, two datasets for 10.5 μM tubulin ± 5 μM Stathmin are reported with solid and dashed lines, and one dataset for 7 μM tubulin ± 2 μM Stathmin in dotted line. Shaded regions represent 95% confidence interval (n±2n) from each dataset in the number of nucleated MTs (n) assuming a Poisson distribution as described in Materials and methods. See Figure 7—figre supplement 1, 2 and Videos 7, 8, 9.

Video 7
XMAP215 increases microtubule nucleation activity of γ-TuRC.

γ-TuRC was immobilized on coverslips and MT nucleation was assayed with low concentration of fluorescent αβ-tubulin (3.5 μM and 7 μM) without (top panels) or with 20 nM XMAP215-GFP (bottom panels). XMAP215 induces MT nucleation from γ-TuRC. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

Video 8
Synergistic microtubule nucleation by γ-TuRC and XMAP215.

Triple-color fluorescence microscopy was performed to observe the molecular sequence of events during MT nucleation from γ-TuRC and XMAP215. γ-TuRC (blue) and XMAP215 (red) formed a complex before MT nucleation occurred (pseudo-colored as green). For 50% of these events, XMAP215 remains on the nucleated minus-end. Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

How does XMAP215 enable MT nucleation from γ-TuRC? We titrated αβ-tubulin at constant γ-TuRC and XMAP215 concentrations and measured the kinetics of nucleation (Figure 7—figure supplement 1C and Figure 7D). XMAP215 effectively decreases the minimal tubulin concentration necessary for MT nucleation from γ-TuRC to 1.6 μM (Figure 7—figure supplement 1C), very close to the minimal concentration for plus-end polymerization. As before, we calculated the composition of the transition state by measuring the power-law dependence between the MT nucleation rate and tubulin concentration with a resulting cooperative assembly of 3.3 ± 0.8 αβ-tubulin dimers occurs (Figure 7E). This suggests that XMAP215 does not lower the thermodynamic barrier to nucleation by altering the geometry of γ-TuRC. Further, neither the N-terminus, containing TOG1-4 domains, nor the C-terminus of XMAP215, containing the TOG5 and C-terminal domain that directly interact with γ-tubulin (Thawani et al., 2018), stimulate additional nucleation from γ-TuRC (Figure 7—figure supplement 1D–E).

Finally, we used our simulations to understand the thermodynamics underlying the MT nucleation activity of XMAP215. Based on its role in accelerating both MT polymerization and nucleation (Thawani et al., 2018; Gunzelmann et al., 2018Flor-Parra et al., 2018), we implicitly modeled the thermodynamic effect of XMAP215’s activity by strengthening the longitudinal tubulin bonds, as described previously (VanBuren et al., 2002). The simulation where only the longitudinal αβ-/αβ-tubulin bond is strengthened does not capture the enhancement of MT nucleation by XMAP215 (Figure 7D, left). Instead, simulations where both the longitudinal γ-/αβ-tubulin and αβ-/αβ-tubulin bond energies are increased by 1.2-fold captures the accelerated kinetics of MT nucleation at low αβ-tubulin concentrations. These simulations also predict a similar transition state composition as measured experimentally (Figure 7D–E, left), supporting XMAP215’s role in strengthening γ-/αβ-tubulin interactions at the nucleation interface. Altogether, our results confirm that XMAP215 indeed functions synergistically with γ-TuRC, in agreement with recent works (Consolati et al., 2020; Thawani et al., 2018; Gunzelmann et al., 2018). Most importantly, our results show that, while the transition state is defined by γ-TuRC’s conformation, XMAP215 strengthens the longitudinal γ-/αβ-tubulin bond to function as a bona-fide nucleation factor.

Inhibition of γ-TuRC-mediated nucleation by specific microtubule-associated proteins

Finally, we asked whether specific MAPs could have an inhibitory effect on MT nucleation from γ-TuRC. The two most abundant inhibitory MAPs in the cytosol, MCAK and Stathmin function by removing αβ-tubulin dimers from the MT lattice (Hunter et al., 2003; Howard and Hyman, 2007) or sequestering αβ-tubulin dimers (Jourdain et al., 1997; Belmont and Mitchison, 1996), respectively. We find that addition of either sub-endogenous concentration of MCAK, or near-endogenous Stathmin concentration (Figure 7F, Figure 7—figure supplement 2 and Video 9) was sufficient to nearly abolish MT nucleation from all γ-TuRC molecules. Thus, γ-TuRC-mediated nucleation is inhibited by MAPs that inhibit MT polymerization.

Video 9
MCAK and Stathmin inhibit γ-TuRC-mediated microtubule nucleation.

Top panels: γ-TuRC was immobilized on coverslips and MT nucleation was observed upon introducing fluorescent 10.5 μM αβ-tubulin without (left) or with 10 nM MCAK (right). Bottom panels: γ-TuRC was immobilized on coverslips and MT nucleation was observed upon introducing fluorescent 10.5 μM αβ-tubulin without (left) or with 5 μM Stathmin (right). Elapsed time is shown in seconds, where time-point zero represents the start of reaction. Scale bar, 10 μm.

Discussion

Decades after the discovery of MTs, their αβ-tubulin subunits and the identification of γ-TuRC as the universal MT nucleator (Oakley and Oakley, 1989; Moritz et al., 1995; Moritz et al., 1998; Zheng et al., 1995; Keating and Borisy, 2000; Wiese and Zheng, 2000), it has remained poorly understood how MTs are nucleated and how this process is regulated in the cell (Kollman et al., 2011; Roostalu and Surrey, 2017; Tovey and Conduit, 2018). Here, we establish a single-molecule assay to study MT nucleation and combine it with computational modeling to identify the rate-limiting, transition state of γ-TuRC-mediated nucleation. We examine how biochemical features of γ-TuRC contribute to its the nucleation activity and regulation.

New methods and direct measurements developed in this study reconcile several prior observations for γ-TuRC-mediated MT nucleation. First, the nucleation activity of γ-TuRC has been found as variable and often low and similar to spontaneous MT assembly (Moritz et al., 1995; Zheng et al., 1995; Kollman et al., 2015; Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020; Kollman et al., 2011; Roostalu and Surrey, 2017), and γ-TuRC’s requirement in the cell has been debated (Hannak et al., 2002; Roostalu et al., 2015; Rogers et al., 2008; Raff, 2019; Woodruff et al., 2017). Low concentration of γ-TuRC molecules obtained from endogenous purifications and lack of live observation of a growing or capped MT minus-end, which is needed to distinguish between γ-TuRC-mediated and spontaneous nucleation, could affect the assessment of γ-TuRC’s nucleation activity. Second, because of technical challenges in the traditional setup where MTs are nucleated, fixed and spun down onto a coverslip (Moritz et al., 1995; Zheng et al., 1995; Kollman et al., 2010; Kollman et al., 2015; Liu et al., 2020; Choi et al., 2010; Liu et al., 2014; Thawani et al., 2018), variable assessment of the role of accessory factors (Liu et al., 2020; Choi et al., 2010; Liu et al., 2014) has been reported. Here, by developing a high-resolution assay that provides specific live information to visualize MT nucleation events from γ-TuRC and distinguish between non-γ-TuRC nucleated MTs, analyses system to measure its nucleation activity independent of concentration, as well as direct visualization of MAPs bound to γ-TuRC or the MT lattice allows us to unambiguously study γ-TuRC-mediated nucleation and its regulation by MAPs.

While the molecular architecture of γ-TuRC was revealed by recent cryo-EM structures (Kollman et al., 2010; Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020), the dynamics of MT nucleation from γ-TuRC and how it relates to γ-TuRC’s specific biochemical features has remained unknown. By combining biochemical investigation with computational modeling, we show that 4 αβ-tubulin heterodimers on neighboring sites form the critical nucleus, that is the rate-limiting transition state on γ-TuRC. A model, in which γ-TuRC stochastically changes its conformation from an open to closed state, where the latter is stabilized by lateral αβ-tubulin interactions, comprehensively explains our experimental measurements. While native γ-TuRC purified from cytosol was used here, further activated γ-TuRC isolated from MTOCs may result in cooperativity between fewer αβ-tubulin dimers for successful nucleation. Likewise, MT assembly from pre-assembled, blunt seeds, could resemble nucleation from already closed γ-TuRCs. We find that the subsequent transition of the growing MT end from blunt- to tapered one, is not the major, rate-limiting step during nucleation from γ-TuRC. Notably, a parallel work also reported MT nucleation from single, human γ-TuRC molecules recently (Consolati et al., 2020). While the majority of findings agree with our work, 6.7 dimers were required in the critical nucleus and an overall lower activity of γ-TuRC (0.5%) was found (Consolati et al., 2020). Low structural integrity of purified γ-TuRC from incorporation of BFP-tagged GCP2 and a higher ratio of γ-tubulin sub-complexes, or species-specific variation in γ-TuRC properties could explain these differences.

Our simulations further predict that a hypothetical low affinity between γ-/αβ-tubulin (Rice et al., 2019) is insufficient to induce any MT nucleation because αβ-tubulins that bind to γ-TuRC dissociate rapidly. Instead, our biochemical investigation show that the high affinity of γ-/αβ-tubulin interaction increases the dwell time of αβ-tubulin dimers on γ-TuRC and promotes γ-TuRC’s MT nucleation activity, as predicted by our modeling. Finally, this net mechanism is thermodynamically favorable compared to spontaneous MT nucleation as the free energy of longitudinal γ-/αβ-tubulin interactions, 13(ΔGLong,γαβ), exceeds the energy penalty from conformational rearrangement of γ-TuRC, ΔGγTuRCconf. In sum, building on the recent structural work (Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020), our results show that the open γ-TuRC conformation and its transition to a closed one defines γ-TuRC’s nucleation activity and transition state. In the future, it will be important to study how γ-TuRC transitions to a closed conformation with high-resolution structural studies, as well as how other biochemical properties, in addition to those modeled here, govern its nucleation activity. Our single molecule assay, kinetic analyses and computational modeling will be essential to complement and place atomic structures into a mechanism that explains how MT nucleation γ-TuRC occurs and how it is regulated.

Although spatial regulation of MT nucleation is achieved by localizing γ-TuRC to specific MTOCs as shown previously (Kollman et al., 2011; Wiese and Zheng, 2006; Tovey and Conduit, 2018; Petry and Vale, 2015), temporal regulation of MT nucleation had been proposed to occur through activation factors that modify γ-TuRC’s conformation and upregulate its activity (Kollman et al., 2011; Choi et al., 2010; Liu et al., 2014; Tovey and Conduit, 2018; Petry and Vale, 2015). While several putative activation factors do not significantly enhance of γ-TuRC’s nucleation activity as shown here, new factors, that are yet to be identified, may serve this role to alter γ-TuRC’s conformation at MTOCs. Alternatively, we postulate another mechanism for temporal control governing the availability and localization of αβ-tubulin. In this model, locally concentrating soluble αβ-tubulin could upregulate the levels of γ-TuRC-mediated MT nucleation, for example as recently shown through accumulation of high concentration of tubulin dimers at the centrosome by MAPs (Woodruff et al., 2017; Baumgart et al., 2019) and by co-condensation of tubulin on MTs by TPX2 during branching MT nucleation (King and Petry, 2020), and finally via specific recruitment of tubulin on γ-TuRC through the binding of XMAP215 as shown here (Thawani et al., 2018; Gunzelmann et al., 2018; Flor-Parra et al., 2018).

Supplementary materials

Supplementary Materials includes nine figures, nine videos, MATLAB code for simulations and source data.

Materials and methods

Purification of recombinant proteins

Request a detailed protocol

Full-length TPX2 with N-terminal Strep II-6xHis-GFP-TEV site tags was cloned into pST50Tr-STRHISNDHFR (pST50) vector (Tan et al., 2005) using Gibson Assembly (New England Biolabs). N-terminal 6xHis-tagged, Xenopus laevis Stathmin 1A was a gift from Christiane Wiese (University of Madison). N-terminal tagged 6xHis-TEV MCAK plasmid was a gift from Ohi et al., 2004. Wild-type XMAP215 with C-terminal GFP-7xHis plasmid was a gift from Reber et al., 2013 and was used to clone XMAP215 with C-terminal SNAP-TEV-7xHis-StrepII tags as well as with C-terminal TEV-GFP-7xHis-StrepII tags, first into pST50 vector and further into pFastBac1 vector. TOG5-CT truncation of XMAP215 was produced by cloning amino acids 1091–2065 into pST50 vector with C-terminal GFP-7xHis-Strep tags. Human γ-tubulin TEV-Strep II-6xHis tags was codon-optimized for Sf9 expression, synthesized (Genscript), and further cloned into pFastBac1 vector. 6xHis tagged γ-TuNA (N-terminal aa 56–89 of Xenopus laevis CDK5RAP2) was also cloned into pST50 and expressed in E. coli Rosetta2 cells. Dual StrepII-6xHis-tagged Xenopus laevis NME7 was cloned into pFastBac1 vector, expressed, and purified from Sf9 cells.

TPX2, Stathmin and truncations of XMAP215 (TOG5-CT and TOG1-4) used in this study were expressed in E. coli Rosetta2 cells (EMD Millipore) by inducing with 0.5–1 mM IPTG for 12–18 hr at 16°C or 7 hr at 25°C. Wild-type XMAP215, MCAK and γ-tubulin were expressed and purified from Sf9 cells using Bac-to-Bac system (Invitrogen). The cells were lysed (EmulsiFlex, Avestin) and E. coli lysate was clarified by centrifugation at 13,000 rpm in Fiberlite F21-8 rotor (ThermoFisher) and Sf9 cell lysate at 50,000 rpm in Ti70 rotor (Beckman Coulter) for 30–45 min.

TPX2 was first affinity purified using Ni-NTA beads in binding buffer (50 mM Tris-HCl pH 8.0, 750 mM NaCl, 15 mM Imidazole, 2.5 mM PMSF, 6 mM BME) and eluted with 200 mM Imidazole. All proteins were pooled and diluted four-fold to 200 mM final NaCl. Nucleotides were removed with a Heparin column (HiTrap Heparin HP, GE Healthcare) by binding protein in 250 mM NaCl and isocratic elution in 750 mM NaCl, all solutions prepared in Heparin buffer (50 mM Tris-HCl, pH 8.0, 2.5 mM PMSF, 6 mM BME). Peak fractions were pooled and loaded on to Superdex 200 pg 16/600, and gel filtration was performed in CSF-XB buffer.

XMAP215-GFP-7xHis was purified using His-affinity (His-Trap, GE Healthcare) by binding in buffer (50 mM NaPO4, 500 mM NaCl, 20 mM Imidazole, pH 8.0) and eluting in 500 mM Imidazole. Peak fractions were pooled and diluted 5-fold with 50 mM Na-MES pH 6.6, bound to a cation-exchange column (Mono S 10/100 GL, GE Healthcare) with 50 mM MES, 50 mM NaCl, pH 6.6 and eluted with a salt-gradient up to 1M NaCl. Peak fractions were pooled and dialyzed into CSF-XB buffer. XMAP215-SNAP-TEV-7xHis-StrepII or XMAP215-TEV-GFP-7xHis-StrepII was first affinity purified with StrepTrap HP (GE Healthcare) with binding buffer (50 mM NaPO4, 270 mM NaCl, 2 mM MgCl2, 2.5 mM PMSF, 6 mM BME, pH 7.2), eluted with 2.5 mM D-desthiobiotin. Peak fractions were pooled, concentrated and further purification via gel filtration (Superdex 200 10/300 GL) in CSF-XB buffer containing 150 mM KCl. For fluorescent labeling of SNAP-tag in XMAP215-SNAP-TEV-7xHis-StrepII, StrepTrap elution was cation-exchanged (Mono S 10/100 GL), peak fractions pooled and reacted with two-molar excess SNAP-substrate Alexa-488 dye (S9129, NEB) overnight at 4°C, followed by purification via gel filtration (Superdex 200 10/300 GL) in CSF-XB buffer. Approximately 70% labeling efficiency of the SNAP-tag was achieved.

γ-tubulin was purified by binding to HisTrap HP (GE Healthcare) in binding buffer (50 mM KPO4 pH 8.0, 500 mM KCl, 1 mM MgCl2, 10% glycerol, 5 mM Imidazole, 0.25 μM GTP, 5 mM BME, 2.5 mM PMSF), washing first with 50 mM KPO4 pH 8.0, 300 mM KCl, 1 mM MgCl2, 10% glycerol, 25 mM imidazole, 0.25 μM GTP, 5 mM BME), and then with 50 mM K-MES pH 6.6, 500 mM KCl, 5 mM MgCl2, 10% glycerol, 25 mM imidazole, 0.25 μM GTP, 5 mM BME) and eluted in 50 mM K-MES pH 6.6, 500 mM KCl, 5 mM MgCl2, 10% glycerol, 250 mM imidazole, 0.25 μM GTP, 5 mM BME. Peak fractions were further purified with gel filtration (Superdex 200 10/300 GL) in gel filtration buffer (50 mM K-MES pH 6.6, 500 mM KCl, 5 mM MgCl2, 1 mM K-EGTA, 1 μM GTP, 1 mM DTT). For covalent labeling of γ-tubulin with Alexa-568 or Alexa-488 dye, peak gel filtration fractions were pooled and dialyzed into labeling buffer (50 mM KPO4 pH 8.0, 500 mM KCl, 1 mM MgCl2, 2% glycerol, 25 μM GDP, 5 mM BME), reacted with 5 to 20-fold excess of Alexa-568 or Alexa-488 NHS ester (catalog # A20003, A20000, GE Healthcare) for 1 hr at 4°C, and unreacted dye was separated with size exclusion Superdex 200 10/300 GL in gel filtration buffer as above. 7% labeling of γ-tubulin was achieved.

γ-TuNA motif from CDK5RAP2 was purified by binding to Ni-NTA resin in binding buffer buffer (50 mM Tris-HCl pH 8, 500 mM NaCl, 20 mM Imidazole), eluted with 250 mM Imidazole and further purified by gel filtration into storage buffer (50 mM Tris-HCl pH 7.5, 200 mM NaCl). NME7 was purified similar to γ-TuNA by first Ni-NTA affinity followed by size exclusion, as described for γ-TuNA, except with salt concentration of 150 mM NaCl and additional 0.05% Tween-20, and further dialyzed into BRB80 for storage.

MCAK was first affinity purified by binding to His-Trap HP (GE Healthcare) in binding buffer (50 mM NaPO4, 500 mM NaCl, 6 mM BME, 0.1 mM MgATP, 10 mM Imidazole, 1 mM MgCl2, 2.5 mM PMSF, 6 mM BME, pH to 7.5), eluting with 300 mM Imidazole, followed by gel-filtration (Superdex 200 10/300 GL, GE Healthcare) in storage buffer (10 mM K-HEPES pH 7.7, 300 mM KCl, 6 mM BME, 0.1 mM MgATP, 1 mM MgCl2, 10% w/v sucrose).

Stathmin was purified using His-affinity (His-Trap HP, GE Healthcare) by first binding in binding buffer (20 mM NaPO4 pH 8.0, 500 mM NaCl, 30 mM Imidazole, 2.5 mM PMSF, 6 mM BME) and eluting with 300 mM Imidazole, followed by gel filtration (HiLoad 16/600 Superdex, GE Healthcare) into CSF-XB buffer (100 mM KCl, 10 mM K-HEPES, 5 mM K-EGTA, 1 mM MgCl2, 0.1 mM CaCl2, pH 7.7 with 10% w/v sucrose).

All recombinant proteins were flash-frozen and stored at −80°C, and their concentration was determined by analyzing a Coomassie-stained SDS-PAGE against known concentration of BSA (A7906, Sigma).

Bovine brain tubulin was labelled with biotin-, Cy5-, Alexa-488 or Alexa-568 NHS esters (GE Healthcare) as described previously (Thawani et al., 2019).

Purification, biotinylated and fluorescent labeling of γ-TuRC

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Endogenous γ-TuRC was purified from Xenopus egg extracts and labeled with the following steps at 4°C. 7–8 ml of meiotic extract from Xenopus laevis eggs, prepared as described previously (Hannak and Heald, 2006; Murray and Kirschner, 1989), was first diluted 5-fold with CSF-XBg buffer (10 mM K-HEPES, 100 mM KCl, 1 mM MgCl2, 5 mM K-EGTA, 10% w/v sucrose, 1 mM DTT, 1 mM GTP, 10 μg/ml LPC protease inhibitors, pH 7.7), centrifuged to remove large aggregates at 3500 rpm (Thermo Sorvall Legend XTR) for 10 min, and the supernatant filtered sequentially with 1.2 μm and 0.8 μm Cellulose Acetate filters (Whatman) followed by 0.22 μm PES filter (ThermoFisher). γ-TuRC was precipitated by incubating with 6.5% w/v PEG-8000k (Sigma) for 30 min and centrifuged at 17,000 rpm (SS-34 rotor, ThermoScientific) for 20 min. γ-TuRC-rich pellet was resuspended in CSF-XB buffer with 0.05% v/v NP-40 using a mortar and pestle homogenizer, PEG was removed via centrifugation at 136,000 xg for 7 min in TLA100.3 (Beckman Ultracentrifuge), and supernatant was pre-cleared by incubating with Protein A Sepharose beads (GE LifeSciences #17127901) for 20 min. Beads were removed, γ-TuRC was incubated with 4–5 mg of a polyclonal antibody custom-made against C-terminal residues 413–451 of X. laevis γ-tubulin (Genscript) for 2 hr on gentle rotisserie, and further incubated with 1 ml washed Protein A Sepharose bead slurry for 2 hr. γ-TuRC-bound beads were washed sequentially with 30 ml of CSF-XBg buffer, 30 ml of CSF-XBg buffer with 250 mM KCl (high salt wash), 10 ml CSF-XBg buffer with 5 mM ATP (removes heat-shock proteins), and finally 10 ml CSF-XBg buffer before labeling. For biotinylation of γ-TuRC, beads were incubated with 25 μM NHS-PEG4-biotin (A39259, ThermoFisher) in CSF-XBg buffer for 1 hr at 4°C, and unbound biotin was removed by washing with 30 ml CSF-XBg buffer prior to elution step. For combined fluorescent and biotin labeling of γ-TuRC, the wash step after ATP-wash consisted of 10 ml of labelling buffer (10 mM K-HEPES, 100 mM KCl, 1 mM MgCl2, 5 mM K-EGTA, 10% w/v sucrose, 0.5 mM TCEP, 1 mM GTP, 10 μg/ml LPC, pH 7.1) and fluorescent labelling was performed by incubating the beads with 1 μM Alexa-568 C5 Maleimide (A20341, ThermoFisher). Unreacted dye was removed with 10 ml CSF-XBg buffer, beads were incubated with 25 μM NHS-PEG4-biotin (A39259, ThermoFisher) in CSF-XBg buffer for 1 hr at 4°C, and unreacted biotin removed with 30 ml CSF-XBg buffer. Labeled γ-TuRC was eluted by incubating 2–3 ml of γ-tubulin peptide (residues 413–451) at 0.4–0.5 mg/ml in CSF-XBg buffer with beads overnight. After 10–12 hr, γ-TuRC was collected by adding 1–2 ml CSF-XBg buffer to the column, concentrated to 200 μl in 30 k NMWL Amicon concentrator (EMD Millipore) and layered onto a continuous 10–50 w/w % sucrose gradient prepared in a 2.2 ml ultra-clear tube (11 × 34 mm, Beckman Coulter) using a two-step program in Gradient Master 108 machine. Sucrose gradient fractionation of γ-TuRC was performed by centrifugation at 200,000xg in TLS55 rotor (Beckman Coulter) for 3 hr. The gradient was fractionated from the top in 11–12 fractions using wide-bore pipette tips and peak 2–3 fractions were identified by immunoblotting against γ-tubulin with GTU-88 antibody (Sigma). γ-TuRC was concentrated to 80 μl in 30 k NMWL Amicon concentrator (EMD Millipore) and fresh purification was used immediately for single molecule assays. Cryo-preservation of γ-TuRC molecules resulted in loss of ring assembly and activity.

Assessment of γ-TuRC with protein gel, immunoblot and negative stain electron microscopy

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To assess the purity of γ-TuRC, 3–5 μl of purified γ-TuRC was visualized on an SDS-PAGE with SYPRO Ruby stain (ThermoFisher) following the manufacturer’s protocol. Biotinylated subunits of γ-TuRC were assessed by immunoblotting with Streptavidin-conjugated alkaline phosphatase (S921, ThermoFisher). For further conjugation of Alexa-568 dye to γ-TuRC, fluorescently labeled subunits were assessed by visualizing an SDS-PAGE gel with Typhoon FLA 9500 (GE Healthcare) with LPG filter and 100 μm pixel size. γ-TuRC purification was also assessed by visualizing using electron microscopy. 4 μl of peak sucrose gradient fraction of γ-TuRC was pipetted onto CF400-Cu grids (Electron Microscopy Sciences), incubated at room temperature for 60 s and then wicked away. 2% uranyl acetate was applied to the grids for 30 s, wicked away, and the grids were air-dried for 10 min. The grids were imaged using Phillips CM100 TEM microscope at 64,000x magnification.

Preparation of functionalized coverslips

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22 × 22 mm, high precision coverslips (170 ± 5 μm, Carl Zeiss, catalog # 474030-9020-000) were functionalized for single molecule assays based on a recent protocol (Roostalu et al., 2015; Bieling et al., 2010) with specific modifications. Briefly, coverslips were labeled on the surface to be functionalized by scratching ‘C’ on right, bottom corner, placed in Teflon racks, sonicated with 3N NaOH for 30 min, rinsed with water and sonicated in piranha solution (2 parts of 30 w/w % hydrogen peroxide and three parts sulfuric acid) for 45 min. Coverslips were rinsed thrice in water, and all water was removed by spin drying completely in a custom-made spin coater. Pairs of coverslips were made to sandwich 3-glycidyloxypropyl trimethoxysilane (440167, Sigma) on the marked sides, placed in glass petri dishes, and covalent reaction was performed in a lab oven at 75°C for 30 min. Coverslips were incubated for 15 min at room temperature, the sandwiches were separated, incubated in acetone for 15 min, then transferred to fresh acetone and quickly dried under nitrogen stream. Coverslip sandwiches were prepared with a small pile of well mixed HO-PEG-NH2 and 10% biotin-CONH-PEG-NH2 (Rapp Polymere) in glass petri dishes, warmed to 75°C in the lab oven until PEG melts, air bubbles were pressed out and PEG coupling was performed at 75°C overnight. The following day, individual coverslips were separated from sandwiches, sonicated in MilliQ water for 30 min, washed further with water until no foaming is visible, dried with a spin dryer, and stored at 4°C. Functionalized coverslips were used within 1 month of preparation.

Imaging chambers were prepared by first assembling a channel on glass slide with double sided tape strips (Tesa) 5 mm apart, coating the channel with 2 mg/ml PLL(20)-g[3.5]- PEG(2) (SuSOS) in dH2O, incubating for 20 min, rinsing out the unbound PEG molecules with dH2O and drying the glass slide under the nitrogen stream. A piece of functionalized coverslip was cut with the diamond pen and assembled functionalized face down on imaging chamber. The prepared chambers were stored at 4°C and used within a day of assembly.

Microtubule nucleation assay with purified γ-TuRC

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The imaging channel was prepared as follows. First, 5% w/v Pluronic F-127 in dH2O was introduced in the chamber (1 vol = 50 μl) and incubated for 10 min at room temperature. The chamber was washed with 2 vols of assay buffer (80 mM K-PIPES, 1 mM MgCl2, 1 mM EGTA, 30 mM KCl, 0.075% w/v methylcellulose 4000 cp, 1% w/v D-(+)-glucose, 0.02% w/v Brij-35, 5 mM BME, 1 mM GTP) with 0.05 mg/ml κ-casien (casein buffer), followed by 1 vol of 0.5 mg/ml NeutrAvidin (A2666, ThermoFisher) in casein buffer, incubated on a cold block for 3 min, and washed with 2 vols of BRB80 (80 mM K-PIPES, 1 mM MgCl2, 1 mM EGTA pH 6.8). Five-fold dilution of γ-TuRC in BRB80 was introduced in the flow chamber and incubated for 10 min. Unattached γ-TuRC molecules were washed with 1 vol of BRB80.

During the incubations, nucleation mix was prepared containing desired concentration of αβ-tubulin (3.5–21 μM) purified from bovine brain with 5% Cy5-labeled tubulin along with 1 mg/ml BSA (A7906, Sigma) in assay buffer, centrifuged for 12 min in TLA100 (Beckman Coulter) to remove aggregates, a final 0.68 mg/ml glucose oxidase (SERVA, catalog # SE22778), 0.16 mg/ml catalase (Sigma, catalog # SRE0041) was added, and reaction mixture was introduced into the flow chamber containing γ-TuRC.

Total internal reflection fluorescence (TIRF) microscopy and analysis of microtubule nucleation from γ-TuRC

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Nucleation of MTs was visualized with inverted Nikon TiE TIRF microscope using a 100X, 1.49 NA TIRF objective. An objective heater collar was attached (Bioptechs, model 150819–13) and the temperature set-point of 33.5°C was used for experiments. Time-lapse videos were recorded for 10 min at 0.5–1 frame per second using Andor iXon DU-897 camera with EM gain of 300 and exposure time of 50–200 ms each frame. Reference time-point zero (0 s) refers to when the reaction was incubated at 33.5°C on the microscope, and for most reactions, imaging was started within 30 s.

Growth speed of the plus-ends of MTs nucleated by γ-TuRC was measured by generating kymographs in ImageJ. For few specific datasets with notable in-plane drift, an ImageJ plugin, StackReg (Thévenaz et al., 1998), was to correct a minor translational drift before proceeding with the analysis. Region of interest (ROI) for individual MTs were selected and resliced to generate a length-time plot and a line was fit to the growing MT plus-end. The slope of this line represents growth speed. The kinetics of MT nucleation from γ-TuRC was measured as follows. A kymograph was generated for every MT nucleated in the field of view. For most nucleation events, the time of nucleation of the MT was obtained from observing the kymograph and manually recording the initiation time point (see Figure 1C for examples). For MTs where nucleation occurred before the timelapse movie began or where the initiation was not clearly observed in the kymograph, the shortest length of the MT that was clearly visible in the timelapse was measured and measured average growth speed of MTs was used to estimate the time of nucleation. We verified that this procedure accurately estimates the nucleation time for test case MTs where the nucleation event was visible. The measurement of number of MTs (N(t)) nucleated versus time was generated from a manual log containing the nucleation time for all MTs observed in the field of view. To represent the theoretical field-to-field heterogeneity in the number of MTs nucleated, we assumed that binding of γ-TuRC and subsequent nucleation follows a Poisson distribution with mean n MTs and standard deviation n MTs. 95% confidence interval in the nucleation measurements, n±2n is displayed on each nucleation time course.

To calculate the percentage of γ-TuRCs that nucleate a MT, we visualized MT nucleation from Alexa-568 labeled γ-TuRC in the presence of 21 μM tubulin and 100 nM XMAP215, or with 10.5 μM tubulin. We counted the number of labeled γ-TuRC molecules attached in the field of view and counted the number of MTs nucleated specifically from these molecules but excluded spontaneous MT nucleation. For the reaction with 21 μM tubulin and 100 nM XMAP215, we directly measured that 15% of γ-TuRC molecules nucleated a MT. For the reaction with 10.5 μM tubulin, a similar calculation was performed and using measured curves (Figure 2C), we estimated the percentage of γ-TuRC that will nucleate with 21 μM tubulin as 11%.

Power-law analysis of critical nucleus size on γ-TuRC

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We consider the following simplified model to determine the number of αβ-tubulin dimers in the rate-limiting, transition state on γ-TuRC that is the critical nucleus. We consider a total number of γ-TuRC molecules  N0 available to nucleate MTs at a specific αβ-tubulin concentration C. The total number of MTs nucleated N(t) from a total N0 γ-TuRCs is a function of time t. If n tubulin dimers assemble cooperatively on γ-TuRC for a successful MT nucleation, the rate of MT nucleation from γ-TuRC molecules available to nucleated at time t, N0-N(t) reads,

(1) dN(t)dt=knucleate(N0-Nt)Cn

Here, we assume that tubulin does not get significantly depleted over time in the course of our reactions as shown by previous calculations (Zanic, 2016). At the start of the reaction  t=0, no MTs have nucleated Nt=0=0, therefore at early times we assume N0-NtN0 to simplify the calculation of the critical MT nucleus,

(2) dNdt|t0=knucleateN0Cn

Converting into log scale,

(3) ln(dNdt|t0)=n ln(C)+a

To obtain the number of αβ-tubulin dimers in the critical nucleus on γ-TuRC, a straight line was fit to the initial, linear region of each nucleation curve N(t) versus t curve for every tubulin concentration C and the rate of nucleation dNdt|t0 was obtained from slope of this fit. A straight line was then fit to ln(dNdt|t0) versus lnC for all concentrations, the slope of which provides the size of critical nucleus n. Finally, the measured rate of nucleation depends on the total number of γ-TuRC molecules available. As the total number of γ-TuRC molecules obtained from different days purifications changes, the rate of nucleation from γ-TuRCs at 10.5μM tubulin was set to 1 (normalization factor) to allow pooling of all datasets for γ-TuRC-mediated nucleation.

Spontaneous microtubule nucleation and data analysis

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Spontaneous MT assembly was visualized similar to γ-TuRC-mediated nucleation with the following changes. The Pluronic, casein and NeutrAvidin incubations were performed identical to γ-TuRC nucleation assay but instead of attaching γ-TuRCs, sucrose-based buffer (of the same composition as used for γ-TuRC elution) was diluted 5-fold with BRB80, introduced in the flow chamber and incubated for 10 min. Washes were performed with 1 vol of BRB80, nucleation mix was added, and imaging was performed as described above. MTs nucleate spontaneously in solution fall down on the coverslip due to depletion forces during the 10 min of visualizing the reaction. The number of MTs nucleated in the field of view were counted manually and plotted in Figure 3B. 95% confidence interval is displayed assuming a Poisson distribution for theoretical field-to-field heterogeneity as described above.

In the absence of any attached nucleation site, the spontaneously nucleated MTs are usually not visualized from the time of their nucleation and the analysis used for γ-TuRC mediated nucleation was adapted. Integrating the Equation (2) above

(4) N(t)=knucleateN0Cnt

Converting into log scale at time t=τ,

(5) lnN(t=τ)=n lnC+b

To obtain the number of αβ-tubulin dimers in the critical nucleus in spontaneous assembly, the number of MTs at a specified time t =7.5 min was measured, a straight line was then fit to lnN(t=τ) versus lnC for all concentrations, the slope of which provides the size of critical nucleus n. All datasets were pooled and reported.

Preparation, microtubule assembly from blunt microtubule seeds and data analysis

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Blunt MTs were prepared with GMPCPP nucleotide in two polymerization cycles as described recently (Wieczorek et al., 2015). Briefly, a 50 μl reaction mixture was prepared with 20 μM bovine brain tubulin with 5% Alexa-568 labeled tubulin and 5% biotin-labeled tubulin, 1 mM GMPCPP (Jena Bioscience) in BRB80 buffer, incubated on ice for 5 min, then incubated on 37°C for 30 min to polymerize MTs, and MTs were pelleted by centrifugation at 126,000 xg for 8 min at 30°C in TLA100 (Beckman Coulter). Supernatant was discarded, MTs were resuspended in 80% original volume of BRB80, incubated on ice for 20 min to depolymerize MTs, fresh GMPCPP was added to final 1 mM, incubated on ice for 5 min, a second cycle of polymerization was performed by incubating the mixture at 37°C for 30 min, and MTs were pelleted again by centrifugation. Supernatant was discarded and MTs were resuspended in 200 μl warm BRB80, flash frozen in liquid nitrogen in 5 μl aliquots, stored at −80°C and found to be stable for months. To verify that these MT seeds have blunt ends, frozen aliquots were quickly thawed at 37°C, diluted 20-fold with warm BRB80, and incubated at room temperature for 30 min to ensure blunt ends as described previously (Wieczorek et al., 2015). MTs were pipetted onto CF400-Cu grids (Electron Microscopy Sciences), incubated at room temperature for 60 s and then wicked away. 2% uranyl acetate was applied to the grids for 30 s, wicked away, and the grids were air-dried for 10 min. The grids were imaged using Phillips CM100 TEM microscope at 130,000 x magnification and most MT ends were found to be blunt.

To assay MT assembly from blunt MT seeds, MT assembly experiments similar to γ-TuRC nucleation assays were performed with the following variation. A lower concentration 0.05 mg/ml NeutrAvidin (A2666, ThermoFisher) was attached, and washes were performed with warm BRB80 prior to attaching MTs. One aliquot of MT seeds was thawed quickly, diluted to 100-fold with warm BRB80, incubated in the chamber for 5 minutes, unattached seeds were washed with 1 vol of warm BRB80, and the slide was incubated at room temperature for 30 min to ensure blunt MT ends. Wide bore pipette tips were used for handling MT seeds to minimize the shear forces that may result in breakage of MTs. Nucleation mix was prepared as described above and a low αβ-tubulin concentration (1.4-8.7 μM) was used. MT assembly from blunt seeds was observed immediately after incubating the slide on the objective heater. Imaging and analysis were performed as described above for to γ-TuRC nucleation assays. The probability curves p(t) for MT assembly were obtained by normalizing for the total number of seeds observed in the field of view N(t)/N0, which allow for direct comparison across datasets. 95% confidence interval represents the theoretical variation in the number of MTs assembled from seeds across fields of view as described above. Rate of nucleation dpdt|t0 was obtained as the slope of a straight line fit to the initial region of p(t) versus t curve for every tubulin concentration C. Power-law analysis was performed similar to γ-TuRC nucleation assays described above. However, as assembly from seeds occur near minimal tubulin concentration needed for polymerization of the plus end C*, the governing equation reads,

(6) dp(t)dt=knucleate(1-pt)Cn-1(C-C*)

At the start of the reaction  t=0, no MTs have nucleated pt=0=0, therefore at early times we assume 1-pt1 to simplify the calculation of the critical MT nucleus. Converting equation (6) in log scale with these simplifications,

(7) ln(dpdt|t0)=(n1)ln(C)+ln(CC)+a

Critical tubulin concentration for polymerization C* was obtained from the x-intercept of the growth speed curve (C*=1.4μM) as described previously. Finally, observing the total number of MT seeds for assembly allows for direct pooling of all datasets for MT assembly from seeds. From fitting a straight line between ln(dpdt|t0) versus lnC-C* for all concentrations, we found the slope n1, which satisfies the above equation and provides the size of critical nucleus for MT assembly from seeds n 1.

Size exclusion chromatography of γ-tubulin and αβ-tubulin

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Size exclusion chromatography of γ-tubulin and αβ-tubulin was performed as follows at 4°C. Purified, human γ-tubulin was diluted to 300 nM in γ-TB buffer (defined 50 mM K-MES pH6.6, 5 mM MgCl2, 1 mM EGTA, 10 mM thioglycerol, 10 μM GDP) with additional 250 mM KCl, and αβ-tubulin individually diluted to 20 μM or 70 μM with BRB80 buffer. Protein aggregates were pelleted by ultracentrifugation of the proteins individually at 80,000 rpm in TLA 100 (Beckman Coulter) for 15 min. γ-tubulin and αβ-tubulin were mixed in 1:1 vol ratio to achieve final concentrations 150 nM γ-tubulin to 10 μM or 35 μM αβ-tubulin and incubated on ice for 10 min. 500 µl of the mixture was loaded onto Superdex 200 Increase 10/300 column (GE Healthcare). The column was equilibrated with γ-TB buffer containing 90 mM KCl and chromatography was performed in this buffer. For control chromatography runs, equal volume of corresponding buffer was used. Absorbance at 214 nm was recorded. 0.3 ml fractions were collected and alternate fractions eluted between 8.5 ml and 16.6 ml were analyzed via immunoblot against γ-tubulin, αβ-tubulin and StrepII tag on γ-tubulin. Secondary antibody conjugated to 800 nm IRDye (LI-COR) was used and imaged with Odyssey CLx imaging station (LI-COR). High-molecular-weight gel filtration standards (Thyroglobulin, Aldolase and Ovalbumin) were purchased from GE Healthcare (Catalog #28403842) and used to estimate the Stokes’ radii of eluted proteins in the same buffer as used for corresponding SEC run (Le Maire et al., 1986).

Measurement of affinity between γ-tubulin and αβ-tubulin with single molecule microscopy

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γ-TuRC nucleation assay was adapted as follows to measure the interaction affinity between γ-tubulin and αβ-tubulin. The imaging channel was prepared by sequentially with 5% w/v Pluronic F-127 incubation, casein buffer washes, 0.05 mg/ml NeutrAvidin incubation in casein buffer, BRB80 washes as described above. 100–200 nM of biotinylated αβ-tubulin or BRB80 buffer was introduced in the flow chamber and incubated for 5 min on a cold block, and unbound αβ-tubulin was washed with 50 μl of BRB80. During the incubations, binding mix was prepared with 100 nM of Alexa-568 or Alexa-488 labeled αβ-tubulin (24–50% labeling percentage) or with 100 nM of labelled γ-tubulin with identical fluorophore (7% labeling percentage) in 1x assay buffer, ultracentrifuged for 12 min in TLA100, oxygen scavengers were added, and reaction mixture was introduced into the flow chamber.

Single molecule binding of fluorescent γ-tubulin or αβ-tubulin with biotinylated αβ-tubulin was visualized with TIRF microscopy using the setup described above at 33.5°C. Images were collected at 2–5 fps with EMCCD gain of 300 and exposure time of 200 ms each frame, and data acquisition was started within 60–90 s after flowing fluorescent γ-tubulin or αβ-tubulin. Minimal photobleaching was observed for the first 15 s of time series acquired, which was used to extract the number by molecules bound by analyzing with the single molecule analysis software ThunderSTORM (Ovesný et al., 2014). Specifically, images were filtered with wavelet B-spline filter (scale 2–3 and order 3), molecules localized with 8-connected local maximum approach, threshold selected as the standard deviation of the first wavelet level, and suggested settings for sub-pixel localization by fitting an integrated Gaussian PSF model with maximum likelihood estimation was performed. The number of single molecules identified for each frame were recorded. The results from ThunderSTORM analysis were verified against manually identified molecules with a sample dataset. To obtain how many molecules bind to biotin-αβ-tubulin for every frame, the number of molecules of γ-tubulin or αβ-tubulin bound inspecifically to the coverslip were independently subtracted from the number of molecules bound to biotin-αβ-tubulin, and this value was divided by the known fluorescent labelling percentage. The calculated number of γ-tubulin or αβ-tubulin bound were averaged for the first 14 s (28 frames) for each dataset, and their mean and standard deviation was reported.

Interaction assays between αβ-tubulin and γ-tubulin were confirmed with biolayer interferometry using Octet RED96e (ForteBio) instrument in an eight-well plate. The plate temperature was held at 33°C and the protein samples were shaken at 400 rpm during the experiment. First, Streptavidin coated biosensors (ForteBio) were rinsed in interaction buffer (50 mM K-MES pH 6.6, 100 mM KCl, 5 mM MgCl2, 1 mM EGTA, 0.05% Tween20, 1 mM GTP). 100–400 nM biotin-labeled αβ-tubulin, or blank buffer, was bound to Streptavidin sensor until loaded protein results in a wavelength shift (Δλ) of 3 nm. Unbound protein was removed by rinsing the sensor in interaction buffer, and interaction with αβ-tubulin was measured by incubating the sensor containing biotinylated αβ-tubulin with 0–35 μM unlabeled αβ-tubulin or 0–1 μM unlabeled γ-tubulin in interaction buffer for 5 min. Δλ (nm) was recorded as a measure of the amount of unlabeled αβ-tubulin that binds to the sensor.

Nucleation of microtubules from purified γ-tubulin

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MT assembly experiments from purified γ-tubulin was performed similar to γ-TuRC nucleation assays described above with following variation. No avidin was attached to the coverslips, and varying concentration of γ-tubulin was prepared by diluting purified γ-tubulin in a high salt buffer (50 mM K-MES pH 6.6, 500 mM KCl, 5 mM MgCl2, 1 mM EGTA), centrifuging to remove aggregates separately for 12 min in TLA100 before adding to the nucleation mix containing 15 μM αβ-tubulin (5% Cy5-labeled) with BSA, glucose oxidase and catalase as described above to a final salt concentration of 44 mM KCl. The reaction mixture was introduced into the flow chamber and imaged via TIRF microscopy. A large number of MTs get nucleated immediately in the presence of 250 nM-1000 nM γ-tubulin.

Negative stain electron microscopy of γ-tubulin filaments

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Purified γ-tubulin was observed to form higher order oligomers previously using analytical gel filtration (Thawani et al., 2018). γ-tubulin filaments were prepared by diluting pure γ-tubulin to 1 μM to the buffer 50 mM K-MES pH 6.6, 5 mM MgCl2, 1 mM EGTA, 100 mM KCl. 5 μl of γ-tubulin mixture was pipetted onto EM grids (Electron Microscopy Sciences, Catalog number: CF400-Cu), which were glow discharged for 25 s. 5 μl sample was incubated on the grid at room temperature for 60 s and wicked away with Whatman filter paper. Grids were washed with 5 μl of dH2O 3 times, stained three times with 0.75% Uranyl formate, where the first two incubations were wicked away immediately while the last was incubated for 30 s. The grids were air-dried for 10 min. Data were collected on a Talos L120C TEM (FEI) equipped with a BM Ceta CCD camera, at a nominal magnification of 74,000x corresponding to a pixel size of 2.03 Å/pixel on the specimen with 1 s integration time, and a defocus range of 1–2 μm underfocus. Micrographs were acquired both in-plane with +0 degree tilt.

Micrographs were converted to mrc file format with IMOD package and imported into RELION-3.0.6 (Punjani et al., 2017) where the data analysis was performed. Contrast transfer function (CTF) estimation of 370 micrographs performed using Gctf (Zhang, 2016). Segments along the length of thin filaments were picked manually. Filaments were boxed into helical segments with 50 Å rise, and subjected to two rounds of 2D classification and particle selection. 1001 particles were selected and were used to generate an ab-initio 3D model. One round of refinement using 3D auto-refine was performed with all particles, followed by one round of 3D classification. 659 particles from the most populated 3D class were selected and another round of refinement was performed to generate a final map with the solvent mask. Analysis was performed in UCSF Chimera (Pettersen et al., 2004). Longitudinal arrangement of αβ-tubulins (pink filament, Figure 4—figure supplement 2E) was generated by isolating one protofilament from PDB: 6DPU (Zhang and Nogales, 2018) and elongating the protofilament with the super-position function in Coot. Lateral arrangement of γ-tubulin array (blue filament, Figure 4—figure supplement 2E) was generated from the crystal contacts observed in the published P21 crystal array (PDB: 1Z5W [Aldaz et al., 2005b]), as described previously (Aldaz et al., 2005a). An alternate γ-tubulin arrangement was also generated by isolating the other possible filament from this P21 symmetry group, where neighboring γ-tubulins neither arrange linearly nor show lateral contacts (green filament, Figure 4—figure supplement 2E). Simultaneous docking of four copies each of longitudinal αβ-tubulin array, lateral γ-tubulin array, or alternate arrangement of γ-tubulin array, was performed by fitting each copy at 15 Å resolution in UCSF Chimera using the fitmap function. Lateral γ-tubulin arrays, but not other filament arrangements, display good fit where the γ-tubulin spacing closely matches that of the reconstructed filaments.

Monte Carlo simulations of microtubule nucleation by γ-TuRC

Simulation procedure

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Kinetic Monte Carlo simulations for MT nucleation and assembly by γ-TuRC were coded and run in MATLAB and were based on a previous stochastic model for the plus-end dynamics of a MT (VanBuren et al., 2002). A type-B MT lattice geometry with 13-protofilaments and a pitch of 3 tubulins at the seam was assumed, and a similar γ-TuRC geometry was encoded. On the blunt plus-end geometry, αβ-tubulin dimers in the MT lattice may have no neighbors, one or half a neighbor at the seam. Once the MT growth occurs into a tapered one, αβ-tubulin dimers can also have one or two neighbors.

New αβ-tubulin dimers arrive with a constant on rate, kon (M−1s−1) on each protofilament. This on rate is equal for each protofilament on the plus-end or on γ-TuRC and remains constant during the simulation. An input concentration of αβ-tubulin dimers was assumed to be constant and not be depleted as shown by previous calculations (Zanic, 2016). Therefore, the net on-rate at each time step is, konC (s-1), where C is the concentration of αβ-tubulin dimers. The interactions between αβ-tubulins was assumed to occur with longitudinal and lateral bond energies, GLong,αβ-αβ and GLat,αβ-αβ, respectively. All αβ-tubulin dimers recruited to the MT lattice or γ-TuRC have a longitudinal bond, and the lateral bond energy depends on the arrangement of neighboring αβ-tubulin dimers. The longitudinal bond energy between γ-/αβ-tubulin on γ-TuRC is GLong,γ-αβ. As a result, the dissociation rate (off-rate) of individual tubulin dimers from the lattice differs and is a function of total bond energy Gtot. Gtot is a sum of the longitudinal bond energy, GLong,αβ-αβ or GLong,γ-αβ, plus the total lateral bond energy from all the neighbors, m×GLat,αβ-αβ. Based on previous works, we also posit that when a tubulin dimer dissociates, all dimers above it in the protofilament dissociate as well. The off-rate of each dimer was then calculated from the following equation as derived previously (VanBuren et al., 2002),

(8) lnK=lnkonkoff (s-1)=-GtotkBT

An open conformation of native γ-TuRC was assumed as observed in recent cryo-EM structures (Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2020). The αβ-tubulins assembled on neighboring sites do not form lateral interactions in the open conformation. A possible transition to a closed γ-TuRC state was allowed with a thermodynamic penalty of ΔGγTuRCconf. However, if n lateral bonds form upon this transition from αβ-tubulins assembled on neighboring sites, that net energy for an open-to-closed transition is ΔGclose=ΔGγTuRCconfnΔGLat,αβαβ. At each time step in the simulation, the rate of this transition is calculated as, kγTuRC-conf ×exp-GclosekBT, where kγTuRC-conf s-1 is the pre-factor of the Arrhenius equation. Hydrolysis of incorporated tubulin dimers was ignored because few catastrophe events were observed in our experiments.

To execute the stochastic simulations, we formulate a list of possible events at every time step, including association of a αβ-tubulin dimer, dissociation of a αβ-tubulin dimer, or transition of γ-TuRC to closed state. The forward rate of each event is calculated as described above. A uniform random number (Ri) from 0 to 1 is generated for each possible event in the list and a single realization of the exponentially distributed time is obtained for each event,

(9) ti=-ln(Ri)ki (s-1)

The event with the shortest execution time is implemented and time elapsed during the simulation is advanced by ti seconds. Each simulation was run with a maximum defined time, usually between 100 and 500 s, or were stopped once the MT grew a total of 2-5 μm in length. The MATLAB code for simulations is provided in the Supplementary Materials.

Parameter estimation

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MT growth parameters were determined by fitting to experimental growth speed curves. Briefly, 20 simulations were performed for each concentration from 2 to 20 μM tubulin for 100 s each. MT length versus time was plotted. Growth speed was obtained from the slope of a linear curve fit of the polymerizing stretch of the length versus time plot. Parameter values of kon=1.3×106 (μM1s1), ΔGLong,αβαβ=7.2kBT, ΔGLat,αβαβ=6.5kBT resulted in the best fit for all tubulin concentrations. These parameter values are similar to those obtained in previous reports (Mickolajczyk et al., 2019; VanBuren et al., 2002). With these polymerization parameters fixed, we varied the remaining parameters. ΔGLong,γαβ was varied from (0.71.3×ΔGLong,αβαβ. GγTuRC-conf was varied from +(0-30)kBT and kγTuRC-conf from (1-0.001) s-1. For each parameter set, we performed 200-500 simulations each at specific tubulin concentrations between 2 and 50 μM. For each simulation, the time of γ-TuRC ring closure was recorded as the nucleation time as it represents the transition from zero MT length to a continuously growing MT. For the simulation where no MT nucleation occurred, a nucleation time of infinity was recorded. Cumulative probability distribution of nucleation (p(t)) versus time was generated from the log of nucleation times for each tubulin concentration. Rate of nucleation dpdt|t0 was obtained by a linear fit from the initial part of each nucleation fit, as described above. The slope of a straight line was fit to ln(dpdt|t0) versus lnC, as outlined in Equation 2,3 above, provide the size of critical nucleus n. The nucleation curves and power-law analysis was compared with experimental data for γ-TuRC-mediated nucleation. The best agreement was found with ΔGLong,γαβ=1.1×ΔGLong,αβαβ, as supported by our biochemical measurements, ΔGγTuRCconf=10kBT and kγTuRC-conf = 0.01 s-1.

To analyze the arrangement of αβ-tubulins in the transition state, the state of γ-TuRC with αβ-tubulin dimers was recorded at the time of γ-TuRC ring closure for 2119 simulations. 3D-dimensional probability distribution of total number of αβ-tubulin dimers and number of lateral αβ-tubulin bonds was generated. The arrangement of αβ-tubulin dimers in the most frequently occurring transition states were displayed with schematics.

To capture the dynamics of MT assembly from blunt seeds, we simulated nucleation assuming a closed γ-TuRC geometry as follows. Lateral bonds between αβ-tubulins assembled on the neighboring sites on γ-TuRC were allowed and GLong,γ-αβ was set equal to GLong,αβ-αβ. Simulations were performed as described above with the following change. The time when the MT in each simulation grew to 50 nm length was recorded to generate the probability distribution. Nucleation curves and power-law analysis was compared with experimental data for seed-mediated MT assembly.

Measuring the effect of microtubule associated proteins on γ-TuRC-mediated nucleation

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Effect of microtubule associated proteins (MAPs) was measured on γ-TuRC’s nucleation activity. γ-TuRC was attached on the coverslips using the setup described above and a control experiment was performed with identical reaction conditions for each protein tested. Because CDK5RAP2’s γ-TuNA motif and NME7 bind γ-TuRC, to test their activity γ-TuRC was additionally incubated 6 μM γ-TuNA motif or 6 μM NME7 to γ-TuRC for 5 min prior to attachment to coverslips to maximize their likelihood of binding and the control γ-TuRC reaction was treated identically with the storage buffer for each protein. Nucleation mix was then prepared containing 10.5 μM αβ-tubulin concentration (5% Cy5-labeled tubulin) as specified along with 1 mg/ml BSA and oxygen scavengers, and either buffer (control), 10 nM GFP-TPX2, 3 μM γ-TuNA motif from CDK5RAP2, 6 μM NME7, 5 μM Stathmin or 10 nM MCAK was added. To test NME7 or MCAK’s effect, the assay buffer additionally contained 1 mM ATP. The reaction mixture containing tubulin and MAP at specified concentration was introduced into the flow chamber containing γ-TuRC, and MT nucleation was visualized by imaging the Cy5-fluorescent channel at 0.5–1 frames per second. For TPX2, fluorescence intensity of the protein was simultaneously acquired.

The number of MTs nucleated over time was measured as described above and the effect of protein on γ-TuRC’s nucleation activity was assessed by comparing nucleation curves with and without the MAP. In order to normalize for the total number of γ-TuRC molecules obtained from different purifications and enable pooling results from all datasets, the number of MTs nucleated at a specified time point, mentioned in each figure legend, was set to 1 for γ-TuRC only (no MAP) control reactions. As before the shaded region represents 95% confidence interval (n±2n)  in the number of MTs, n assuming a Poisson distribution that determines binding and subsequent nucleation from γ-TuRCs and was calculated and displayed on each nucleation time-course.

Cooperative microtubule nucleation assay with purified XMAP215 and γ-TuRC

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A similar set of experiments as above to characterize the effect of MAPs was performed to study the effect of XMAP215 on γ-TuRC-mediated nucleation with the single molecule assays with the following differences. 20 nM of XMAP215-GFP-7xHis was added to nucleation mix prepared with 3.5–7 μM αβ-tubulin concentration (5% Cy5-label) in XMAP assay buffer (80 mM K-PIPES, 1 mM MgCl2, 1 mM EGTA, 30 mM KCl, 0.075% w/v methylcellulose 4000 cp, 1% w/v D-(+)-glucose, 0.007% w/v Brij-35, 5 mM BME, 1 mM GTP). MTs nucleated from attached γ-TuRC with and without XMAP215 were measured to assess the efficiency of nucleation induced by XMAP215. To assess if N- or C-terminal domains of XMAP215 increases nucleation efficiency, wild-type XMAP215 was replaced with a C-terminal construct of XMAP215 (TOG5-Cterminus-GFP) or an N-terminal construct (TOGs1-4-GFP) in the described experiment.

To measure the kinetics of cooperative nucleation XMAP215 and γ-TuRC, a constant density of γ-TuRC was attached as described above and nucleation mix nucleation mix was prepared with a range of αβ-tubulin concentration between 1.6 and 7 μM (5% Cy5-label) with 20–25 nM of XMAP215-GFP-7xHis or XMAP215-TEV-GFP-7xHis-StrepII in XMAP assay buffer, introduced into reaction chamber and MT nucleation was imaged immediately by capturing dual color images of XMAP215 and tubulin intensity at 0.5 frames per second.

Data analysis was performed as above for γ-TuRC mediated nucleation, theoretical field-to-field heterogeneity in the number of MTs nucleated was represented with a Poisson distribution as before and 95% confidence interval. Critical tubulin nucleus for cooperative nucleation from XMAP215 and γ-TuRC was obtained as described for γ-TuRC alone (Equations 1, 2, 3). A straight line was fit to log rate of nucleation ln(dNdt|t0) versus log tubulin concentration lnC and its slope provides the size of critical nucleus n. Finally, to normalize for the total number of γ-TuRC molecules obtained from different purifications, the rate of cooperative nucleation from XMAP215 and γ-TuRC at 3.5μM tubulin was set to 1. All datasets were pooled and reported.

Triple-color imaging of XMAP215, γ-TuRC and microtubules

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For triple-color fluorescence assays, Alexa-568 and biotin-conjugated γ-TuRC was first attached to coverslips as described above with the following variation: 0.05 mg/ml of NeutrAvidin was used for attaching γ-TuRC. Nucleation mix was prepared with 7 μM αβ-tubulin (5% Cy5-label), 10 nM Alexa-488 XMAP215-SNAP or XMAP215-GFP with BSA and oxygen scavengers in XMAP assay buffer (80 mM K-PIPES, 1 mM MgCl2, 1 mM EGTA, 30 mM KCl, 0.075% w/v methylcellulose 4000 cp, 1% w/v D-(+)-glucose, 0.007% w/v Brij-35, 5 mM BME, 1 mM GTP) and introduced into the reaction chamber containing attached γ-TuRC. Three-color imaging per frame was performed with sequential 488, 568 and 647 nm excitation and images were acquired with EMCCD camera at 0.3 frames per second.

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Decision letter

  1. Jens Lüders
    Reviewing Editor; Institute for Research in Biomedicine, Spain
  2. Anna Akhmanova
    Senior Editor; Utrecht University, Netherlands
  3. Jens Lüders
    Reviewer; Institute for Research in Biomedicine, Spain

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This study establishes TIRF-based single molecule imaging of microtubule nucleation from immobilized, purified γ-TuRCs, which function as nucleation templates. The authors provide first insight into the molecular events that occur at the γ-TuRC during nucleation and establish the critical size that the α-β-tubulin nucleus requires to be converted into a growing microtubule.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Molecular mechanism of microtubule nucleation from γ-tubulin ring complex" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Jens Lüders as the Reviewing Editor and Reviewer #3, and the evaluation has been overseen by a Senior Editor.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

Establishing an in vitro assay that allows to study microtubule nucleation events from purified gTuRC at the single molecule level is overdue. Such an assay is required to answer fundamental and long-standing questions regarding the mechanism of nucleation. In this regard the assay using immobilized gTuRC and single molecule TIRF imaging, as presented in the current manuscript, is an important achievement. Unfortunately, it has not yet been used to its full potential. Several interesting observations are made, but the chosen path and derived conclusions in this manuscript are challenged by three recently published Cryo-EM structures of native gTuRC, including gTuRC purified from Xenopus egg extract as in the present study (Liu et al., 2019). The structures show that native gTuRC does not match the symmetry of a microtubule. It is in an 'inactive' conformation, one that is not well suited to promote lateral tubulin interactions, questioning the validity of the presented model. Considering this and other issues raised by the reviewers the manuscript does not provide the conceptual advance required by eLife.

Reviewer #1:

The authors use γ-TuRCs purified from Xenopus egg extracts in combination with single molecule microtubule nucleation assays to gain insight into the process of microtubule nucleation. The key findings are that γ-TuRCs enhance microtubule nucleation by promoting the lateral association of a/β-tubulin molecules and that nucleation from γ-TuRCs is further promoted by XMAP-215. The experiments are well-executed and the data is nicely presented (although I have some questions about N numbers – see below). Overall, I think there are some interesting insights into γ-TuRC-mediated nucleation and the experiments are at the cutting edge, but I feel that we do not learn a great deal more about the role of γ-TuRC and XMAP-215 in microtubule nucleation to justify publication within a broad spectrum high-impact journal such as eLife. Several results or notions are already known e.g. γ-TuRCs and XMAP-215 promote microtubule nucleation, a higher concentration of tubulin is required for nucleation than for microtubule elongation, γ-tubulin forms filaments, XMAP-215 promotes nucleation, XMAP-215 associates with γ-TuRCs. Moreover, the main message, that γ-TuRCs promote lateral association of tubulin dimers, is widely assumed based on γ-TuRC structure and because longitudinal a/β-tubulin contacts are believed to be stronger than lateral contacts. As the authors themselves state, "the insights on MT nucleation by γ-TuRC and XMAP215 provide an essential basis to build upon". I therefore feel that, without further advances, such as how γ-TuRCs can be activated to better promote microtubule nucleation, the paper would be better suited to a more specialised journal, such as JBC.

More specific comments:

1) The title should be more specific to the work done, as the current title indicates that the authors have solved the entire mechanism of nucleation.

2) Abstract: "the underlying mechanism largely remains a mystery". This is a difficult statement to swallow. I would argue that the underlying mechanism (templating the addition of tubulin dimers into a tube-like structure) has been widely assumed (although not directly proved) for a long time, in particular after the 2010 Nature paper from David Agard's group that showed the structure of the yeast "γ-TuRC". This was good proof that the template model was correct, rather than the protofilament model. I think the authors should therefore modify their statement to something like: "While it has been assumed that…., here we provide direct evidence for….".

) Abstract: "…we uncover that γ-TuRC nucleates a MT more efficiently than spontaneous assembly". This was already known – many studies have shown (using more standard microtubule nucleation assays) that microtubules form more readily in the presence of γ-TuRC.

4) Figure 1B: Three γ-TuRC nucleation sites are indicated by arrows – is this all of the γ-TuRCs present within this field (Can the authors show a two-colour image overlay)? If there were only three γ-TuRCs, it would not fit with the finding that spontaneous nucleation does not occur at 10.5μM tubulin….

5) Subsection “Defining the microtubule nucleus on γ-TuRC”: "Surprisingly, γ-TuRC nucleated MTs starting from 7 μM tubulin (Figure 1D), which is higher than the minimum tubulin concentration (C*) needed for growth at pre-formed MT plus-ends (C* = 1.4μM, Figure 1E)". I don't think this is surprising, given the results of Wieczorek et al., 2015 – these authors used solution exchange to show that, while nucleation from a microtubule seed required ~6μM tubulin, microtubules could continue to grow at ~1μM tubulin. This statement should be toned down and the authors should refer to Wieczorek et al. results.

6) Figure 1G: The authors use power-law calculations to calculate that ~3.7 tubulin dimers are required for nucleation. It is not clear exactly how they did this. Given that this power analysis underlies the main finding of the paper, I think it deserves a full explanation.

7) Subsection “Does γ-TuRC nucleate a microtubule via the blunt plus-end model?”: The authors use microtubule seeds to compare their nucleation efficiency to γ-TuRCs and find that microtubules can assemble from these seeds at only 2.45μM tubulin. This is significantly lower than that found by Wieczorek et al., 2015 (6μM tubulin) – can the authors comment on this and suggest an explanation?

8) Subsection “Molecular insight into microtubule nucleation by γ-TuRC”: Experiments to nucleate microtubules from γ-tubulin oligomers: I don't think this adds much to the paper. It is already known that γ-tubulin can form filaments and it is hard to understand how these filaments would nucleate microtubules, given that the filaments are presumably made of laterally contacting γ-tubulin molecules, thus creating a linear, rather than circular, template for the addition of a/β-tubulin dimers. It was also not clear from the methods what the final salt concentration was during the nucleation reactions, given that the g-tubulin was diluted in high salt buffer before being added to the reaction buffer. This is important as the authors show that tubulin oligomers do not form at high salt concentrations. If the authors want to keep these experiments in the paper, they need to cite work from the groups of Alvarado-Kristensson, Binarova and Monasterio who have shown that γ-tubulin can oligomerise into filaments.

9) Subsection “Molecular insight into microtubule nucleation by γ-TuRC”: The stochastic models, used to predict how many γ-tubulin sites must to be occupied to have the best chance of 2 adjacent tubulin dimers being present, are based on a perfectly circular arrangement of γ-tubulin molecules (Figure 2H). Liu et al., 2019 have just shown the cryo-EM structure of the Xenopus γ-TuRC. It is in an open confirmation, where the first and last γ-tubulin molecules are not adjacent to each other. I presume this would change (albeit perhaps only slightly) the calculations from the stochastic model simulations, so the authors should repeat them using an open ring where there is no lateral contact if positions 1 and 13 are occupied.

10) For quantifying the numbers of microtubules nucleated in e.g. Figure 1F, Figure 3A,3C it is not clear how many times the experiments were performed and whether the graphs represent mean or median values. Given the shakiness of the lines in the graph in Figure 3A, the N numbers seem low. I would expect that the experiments (being quick to perform) would have been performed several times (perhaps 5 to 10 times) and that a mean or median value was used to represent microtubule numbers at each timepoint. Can the authors confirm this, or do more repeats?

11) Figure 3A: the authors show that TPX2 has no role in microtubule nucleation from γ-TuRCs, but previous studies (Wieczorek et al., 2015, Roostalu et al., 2015, Woodruff et al., 2017) have shown that TPX2 does promote nucleation. Do the authors believe there is a difference in the requirement of TPX2 when considering γ-TuRCs versus microtubule seeds as templates? Can the authors repeat previous results showing that TPX2 promotes nucleation from microtubule seeds, as a positive control to show that their purified TPX2 is functional?

12) Subsection “How do γ-TuRC and XMAP215 synergistically nucleate microtubules?”: "XMAP215 effectively decreases the minimal tubulin concentration necessary for MT nucleation from γ-TuRC to 1.6 μM". How was this calculated? Also, does adding XMAP215 also increase the probability of spontaneous nucleation? In a model where XMAP215 binds γ-tubulin and uses TOG domains to add tubulin dimers for the initial nucleation event, the addition of XMAP215 should not increase the probability of spontaneous nucleation.

13) Figure S4B: the authors test whether the C-term γ-tubulin-interacting region of XMAP215 promotes microtubule nucleation, and show that it does not. It would be informative to also test whether the rest of XMAP-215 alone (i.e. XMAP-215 minus the γ-tubulin binding region) can promote microtubule nucleation from a γ-TuRC. Presumably not, but this truncated form of XMAP-215 should in theory be able to bind to microtubule plus ends via the TOG domains. It would therefore be a nice experiment to show that it is not plus-end bound XMAP-215 that is promoting nucleation, but it is the XMAP-215 that binds to the γ-TuRC.

Reviewer #2:

The present manuscript uses purified γ-TuRC, tubulin, and different MAPs, to reconstitute microtubule nucleation in vitro, and to quantify nucleation rates, growth speed, and cooperativity of tubulin dimers. Using coated coverslips to anchor tagged γ-TuRCs, in combination with TIRF microscopy, several important observations are made:

- Nucleation of microtubules occurs from γ-TuRCs de novo, and this is more efficient than spontaneous assembly.

- The critical nucleus for nucleation from γ-TuRCs comprises only 3-4 tubulin dimers, which is less than half the size required for spontaneous assembly.

- γ-TuRC-mediated nucleation differs from plus-end growth at blunt microtubule ends, where a single tubulin dimer suffices as a nucleus.

- γ-TuRC-mediated nucleation is not influenced by TPX2. By contrast, XMAP215 decreased the minimal tubulin concentration and increased the nucleation rate, by binding first to the γ-TuRC, yet requiring a similar critical nucleus of 3-4 tubulin dimers.

Altogether, these data provide extremely valuable insight into the mechanisms of microtubule nucleation, since they clarify multiple major issues: γ-TuRCs act directly as templates for microtubule nucleation, ruling out other hypotheses that propose spontaneous nucleation, followed by capping of the free minus-ends by γ-TuRCs. Most importantly, we have for the first time a concrete idea about the critical size of the tubulin nucleus, and about the involvement (or non-involvement) of XMAP215 and TPX2 in the nucleation process.

The manuscript is timely, since several other studies on the γ-TuRC (e.g. structure of the native full complex) are likely to be published by other groups in the very near future.

Reviewer #3:

The manuscript by Thawani et al., together with a recently posted study by the Surrey group on the bioRxiv preprint server, is the first to study microtubule nucleation from purified, immobilized gTuRC at the single molecule level by TIRF microscopy. Using this approach the authors investigate and compare nucleation from gTuRC with spontaneous MT assembly and nucleation from preformed MT seeds. They find that nucleation from gTuRC is more efficient than spontaneous assembly, but less efficient than nucleation from MT seeds. They argue that this is due to relatively weak longitudinal affinity between g-tubulin and ab-tubulin based on measurements of ab-tubulin interaction with purified g-tubulin and ab-tubulin in biolayer interferometry assays. XMAP215 significantly increases gTuRC nucleation activity. They estimate that nucleation from gTuRC alone requires 3-4 ab-tubulin dimers to bind to gTuRC before the kinetic nucleation barrier is overcome and argue that this is facilitated by the lateral array of g-tubulins in the gTuRC.

The establishment of TIRF imaging of nucleation from gTuRC at single molecule level is overdue and the presented work is a great achievement in this regard, providing interesting new ideas and experimental possibilities. However, the authors base their conclusions on various assumptions that cannot be made without further evidence and control experiments. A major issue is the fact that native gTuRC is not a perfect template that matches the symmetry of a microtubule, as revealed by three independent recent Cryo-EM studies by the Kapoor, Schiebel and Surrey labs. Overall, the study requires significant additional work that in my opinion would go beyond what would be reasonable for a revision in eLife.

Major points:

1) Subsection “Reconstituting and visualizing microtubule nucleation from γ-TuRC”: "MT nucleation events occurred specifically from single gTuRC molecules" – gTuRCs are not labelled in this assay, how do the authors get to this conclusion? How can they exclude that some of the MTs are from spontaneous nucleation?

2) The main conclusions of this study regarding the nucleation mechanism are based on the assumption that gTuRC is a perfect template, which has recently been disproven. Cryo-EM data of purified native human and Xenopus gTuRC clearly show that the g-tubulin array in gTuRC does not match the symmetry of a MT, by displaying lateral gaps between g-tubulin molecules, in particular in the part of gTuRC that contains GCP4, 5, and 6. This would be expected to impact its ability to promote lateral interactions between ab-tubulin dimers.

3) Figure 2C-E: It is assumed that γ-tubulin alone, through lateral interaction, forms a filament-like microtubule nucleation template. There are two issues here. First, there is no evidence that the g-tubulin filaments are indeed formed by lateral g-tubulin interactions. In fact, the EM image in S2C shows a fiber much thicker than expected. Second, the authors should present evidence that the formed ab-tubulin polymers are actually microtubules rather than other types of filaments (e.g. protofilaments or polymerization along g-tubulin filaments).

4) Fig2D: Are these really protofilaments? They seem much thicker. Also, can the authors exclude that g-tubulin polymerizes during coating of the chip?

5) Figure 2H: First, the simulation (i) assumes that gTuRC is perfectly ring-shaped (but it is not, see point 2 above) and ii) does not take into account binding affinity with tubulin dimers already bound to the template. After binding of the first dimer, binding of a second dimer at the neighbouring position may be promoted by lateral affinity. Also, I don't understand how the probability in calculated – should it not increase further after 4 sites are occupied? At this point it should be much more likely to have lateral contacts than not.

6) Considering the technical challenges with experimental repetitions and use of different, freshly prepared gTuRC preps for each repetition, it would be useful to repeat the nucleation assay to measure spontaneous and nucleation from seeds and from gTuRC side-by-side to properly compare the different conditions. It should be discussed that the minimal concentration for nucleation from seeds in a previous study by Wieczorek et al., (2015) was quite different and similar to nucleation from gTuRC in the present study.

7) It is not very clear how many times experiments were repeated and what the exact outcomes of each were. Sometimes data from different repetitions was "pooled", sometimes only one "representative dataset" of several repetitions is shown. Would it not be more appropriate to plot mean values from the multiple repetitions? The descriptions in all three figure legends are quite vague, for example Figure 2: "…were repeated at least twice with multiple supporting results…" – what exactly does this mean?

The same is true for the transparent reporting form:.…"confirmation and supporting experiments were performed with slightly different conditions and complete agreement was found amongst all measurements. The datasets were sufficient to give confidence on the measured value of interest…". What are the slightly different conditions? How are "complete agreement" and "confidence" determined?

Source data is not provided because "all data generated in this study was pooled (no data was left out)", but if only one "representative dataset" is shown, my understanding is that some data was in fact left out.

8) What is the percentage of gTuRCs that nucleate microtubules in the TIRF assay? This number should be provided.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "The transition state and regulation of microtubule nucleation from γ-TuRC revealed by single molecule microscopy" for consideration by eLife. The evaluation of your paper has been overseen by a Reviewing Editor and Anna Akhmanova as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.

Summary:

The assembly and organization of the microtubule cytoskeleton requires nucleation of microtubules by the nucleator γ-TuRC, but the underlying mechanism has remained obscure. Recent Cryo-EM structures of γ-TuRC have raised additional questions by showing that native γ-TuRC is in a conformation that is not well suited to template microtubule nucleation. This study provides first insight by establishing TIRF imaging from purified, immobilized γ-TuRCs in vitro. Using a combination of biochemistry and modeling the study explains how an "imperfect" nucleation template can function as nucleation platform and identifies several crucial determinants of this activity. Rather than activation of γ-TuRC by a conformational switch prior to nucleation, the authors propose that the conformational switch occurs as a result of nucleation.

The authors have provided very thorough revisions, presenting and including a substantial amount of new data. The manuscript has been significantly improved and despite the lack of structural support data by EM in the manuscript, the main conclusions are well supported and we consider it now suitable for publication in eLife. There are, however, a few remaining issues that we would like the authors to address.

Essential revisions:

1) We appreciate that you have modified the title, but it now seems to not make complete sense. We would suggest "The transition state and regulation of γ-TuRC-mediated microtubule nucleation revealed by single molecule microscopy".

2) "Notably, the distinct minimal tubulin concentration needed for seeds to assemble a MT as compared to a previous report 24 results from the differences in assay conditions." The authors should be clearer here, explaining in more detail (as they did in the response to reviewers) that they got the same result as previous authors when they used their particular conditions.

3) "In summary, because γ-TuRC positions an array of γ-tubulins at its nucleation interface that are thought to stabilize intrinsically weak, lateral αβ/αβ-tubulin interaction 9,10,13,14,17,20,22147, MT nucleation by γ-TuRC has been proposed to function similar to polymerization of a MT end. Here we show several lines of evidence that γ-TuRC-mediated nucleation has distinct characteristics from MT polymerization and assembly from blunt MT seeds. While growth speed of MTs nucleated from γ-TuRC or templated from MT seeds is similar (Figure 3—figure supplement 1D), γ-TuRC molecules do not nucleate MTs at low tubulin concentration where MT polymerization can occur." It is important to point out here that this relates specifically to the purified γTuRCs that nucleate microtubules in the absence of other cellular factors. While the authors do show that some of the proposed activators do not activate the γ-TuRC in their assays, they cannot yet rule out that other proteins do function in vivo to promote the correct configuration of the γTuRC. They mention this in the Discussion section, but they are making the statement in the Results section, and it is a bit misleading to group all γTuRCs into this poor-nucleation bracket.

4) Figure 6B, D – the authors state that there is no significant difference in nucleation rate when they add γTuNA or TPX2, but the graphs do show differences and no statistical test is provided.

5) "Recently, XMAP215 was discovered to be a nucleation factor that synergizes with γ-TuRC". There is a new pre-print from Trisha Davis' group showing that nucleation by γTuRC and XMAP215 is additive, not synergistic http://biorxiv.org/content/early/2020/05/23/2020.05.21.109561. It would be useful for the reader if the authors would discuss how this compares to their work here.

6) The authors should discuss in more detail recently published work by the Surrey group (2020), which also shows, as the current manuscript, TIRF imaging of nucleation from immobilized γ-TuRCs. There are some differences e.g. regarding the critical number of dimers that need to assemble on gTuRC and regarding overall gTuRC activity in the assay. It would be very useful for the reader to know how the observations/conclusions in the two studies agree or differ.

https://doi.org/10.7554/eLife.54253.sa1

Author response

[Editors’ note: The authors appealed the original decision. What follows is the authors’ response to the first round of review.]

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

Establishing an in vitro assay that allows to study microtubule nucleation events from purified gTuRC at the single molecule level is overdue. Such an assay is required to answer fundamental and long-standing questions regarding the mechanism of nucleation. In this regard the assay using immobilized gTuRC and single molecule TIRF imaging, as presented in the current manuscript, is an important achievement. Unfortunately, it has not yet been used to its full potential. Several interesting observations are made, but the chosen path and derived conclusions in this manuscript are challenged by three recently published Cryo-EM structures of native gTuRC, including gTuRC purified from Xenopus egg extract as in the present study (Liu et al., 2019). The structures show that native gTuRC does not match the symmetry of a microtubule. It is in an 'inactive' conformation, one that is not well suited to promote lateral tubulin interactions, questioning the validity of the presented model. Considering this and other issues raised by the reviewers the manuscript does not provide the conceptual advance required by eLife.

Although all reviewers recognized this work as far-reaching in studying MT nucleation from γ-TuRC at the single molecule level and a necessary step forward, the results from the original manuscript did not account for key information on the structure and conformation of γ-TuRC described in the works that were published in the last months (Liu et al., 2020; Wiezcorek et al., 2020) while our original manuscript was in review. We took this feedback seriously and performed new experiments and computational modeling to address these open questions. We now provide a more comprehensive understanding of how MT nucleation from γ-TuRC occurs as described below and believe that the revised work provides the conceptual advance for the microtubule and the broader cytoskeleton fields.

Re-examination of g-/αβ-tubulin interaction

We re-examined the biochemical interaction between g-tubulin and αβ-tubulin to address the comments raised by reviewer 3. First, we measured the interaction of purified g-tubulin and αβ-tubulin via size exclusion chromatography (SEC) experiments (Figure 4A and Figure 4—figure supplement 1A). We performed SEC of g-tubulin (150nM) with αβ-tubulin at two different concentrations, 10μM (low) and 35μM (high), as well as of g-tubulin and αβ-tubulin alone. At low concentration of αβ-tubulin alone, we observed a single species for αβ-tubulin heterodimer eluting in fractions H-K (Figure 4—figure supplement 1A(i)). At high concentration of αβtubulin alone, we additionally observed αβ-tubulin oligomers in our setup (Figure 4—figure supplement 1A(ii)), red arrows marked in fractions B-C) that correspond to two αβ-/αβ-tubulin in complex from estimated molecular weight using the gel filtration standards. While, g-tubulin alone runs as a broad peak in fractions I-N (Figure 4A(i)), upon incubation with αβ-tubulin, g-tubulin additionally elutes at an earlier elution volume in fraction H ((Figure 4A(ii-iii)) and its elution profile is modified to following αβ-tubulin. Surprisingly, this was observed with both low and high αβ-tubulin concentration, showing that the g-/αβ-tubulin interaction occurs even at low concentrations and suggests that this interaction is higher affinity than that between two αβ-/αβ-tubulins, which requires higher concentrations.

Second, we turned to single molecule binding assays as an orthogonal approach (Figure 4B). We attached buffer or biotin-αβ-tubulin molecules on the coverslip at low concentration of 100-200nM and introduced fluorescent αβ-tubulin (Figure 4B(i)) or fluorescent g-tubulin molecules (Figure 4B(ii)) in solution also at low concentrations (50-100nM) to avoid formation of MTs or g-tubulin assemblies. In both cases, specific binding of αβ-tubulin and g-tubulin molecules binding to biotin-αβ-tubulin on the coverslip was observed (compare left- and right-panels in Figure 4B(i-ii)), yet more fluorescent g-tubulin molecules than fluorescent αβ-tubulin molecules bound. Performing single molecule analysis while accounting for total labelled percentage of αβ- and g-tubulin in our purifications as detailed in the Methods section, revealed that nearly 15-fold more g-tubulin molecules than αβ-tubulin bind to the attached biotin-αβ-tubulin molecules (Figure 4B).

Lastly, we verified this interaction with biolayer interferometry (Figure 4—figure supplement 1B), which corroborated the SEC and single molecule assays. We would like to note that a reverse configuration was used for interferometry assays in the previous version of the manuscript where g-tubulin was attached on the probe via its His-tag and αβ-tubulin was introduced in solution, as compared to the revised version.

Upon further investigation, we found that the proximity of the His-tag, used as the site of attachment to the probe, to the αβ-tubulin’s binding surface occluded the binding of αβ-tubulin, both in the interferometry and single molecule assays (data not shown). Further, g-tubulin shows significant non-specific background binding to the probe in interferometry experiments (Figure 4—figure supplement 1B(ii)). To overcome these drawbacks in the revised manuscript, we developed new binding assays that either do not involve any immobilization of proteins (SEC) or and where attached molecules could be directly visualized (single molecule assay). These biochemical measurements, therefore, serve as most direct assessment of binding and supersede the interferometry experiments now presented in the supplement.

In sum, we find that a high affinity g-/αβ-tubulin interaction at the nucleation interface promotes gTuRC’s MT nucleation activity. We are grateful to reviewer 3 for their feedback that prompted us to assay g-/αβ-tubulin interactions with alternate methods which resulted in a more comprehensive biochemical understanding.

Monte Carlo simulations of microtubule nucleation from γ-TuRC

We next developed a detailed mathematical model to capture the dynamics of MT nucleation from γ-TuRC. Our Monte Carlo simulations were based on previous models for polymerization and depolymerization of the MT plus-end, first developed by Odde and Cassimeris (VanBuren et al., 2002). We assumed a 13-protofilament (pf) geometry of γ-TuRC and the MT lattice with a pitch of 3 tubulin monomers (Figure 5A). The assembly of αβ-tubulin on γ-TuRC or the MT lattice occurred with a constant on-rate kon (μM-1s1pf-1) with longitudinal and lateral Gibbs free energy of interaction with other αβ-tubulins, ∆GLong,αβ-αβ and ∆GLat,αβ-αβ, respectively, and with longitudinal free energy of interaction with gtubulin at the interface of γ-TuRC, ∆Long,γ-αβ. To incorporate the data obtained from recent cryo-EM structures (Liu et al., 2020; Wiezcorek et al., 2020), we assumed that native γ-TuRC has an open conformation where interactions between tubulins on neighboring sites could not occur. A thermodynamic penalty,∆GγTuRC-conf, and a rate constant kγTuRC-conf(s-1) determine the transition from this open to closed γ-TuRC conformation where lateral tubulin interactions can occur. We ran stochastic simulations with these assumptions and determined the input parameters from our experimental data.

We determined the values for parameters kon, ∆GLong,αβ-αβ and ∆GLat,αβ-αβ by fitting our experimental measurements of growth speed (Figure 5—figure supplement 1A). The resulting values are in agreement with previous estimates for these parameters (VanBuren et al., 2002; Mickolajczyk et al., 2019). ∆GLong,γ-αβ was estimated to be 1.1 times ∆GLong,αβ-αβ based on our experimental binding affinity measurements (Figure 4). Finally, we explored the behavior for different values of the two unknown parameters ∆GγTuRC-conf and kγTuRC-conf. We note that the total energy of transition to a closed γ-TuRC state is affected by the number of tubulin dimers that bind on the neighboring sites on the open γ-TuRC. Specifically, the free energy of all (n) lateral contacts that can form upon closure adds to the total transition energy between the start and end state, ∆GγTuRC-conf + nGLat,αβ-αβ.

With this model, we could incorporate features of γ-TuRC that contribute to its nucleation activity including its conformation, blunt-end geometry and differential biochemical interactions. To our surprise, the simulations recapitulate our experimental measurements in detail. First, we observe a sharp transition from zero MT length on γ-TuRC to a continuously growing MT (Figure 5B). Second, this transition occurs at variable time for each γ-TuRC and the resulting nucleation curves obtained from hundreds of simulations match our experimental curves (Figure 5—figure supplement 1B(i)-C(i)). Analogous to our experimental results, we measured the cooperativity of αβ-tubulin dimers in the rate-limiting transition state and found that it is determined by the magnitude of ∆GγTuRC-conf (Figure 5C) but not affected significantly by changing kγTuRC-conf or ∆GLong,αβ-αβ (Figure 5—figure supplement 1B(ii)-C(ii)). For the specific parameter values of ∆GγTuRC-conf = +10kbT and kγTuRC-conf = 0.01s−1, the nucleation kinetics strikingly agree with our experimental data for γ-TuRC-mediated MT nucleation (Figure 5—figure supplement 2A). Hundreds of these simulations do not result in successful MT nucleation at αβ-tubulin concentration less than 7μM, and increasing MT nucleation events occur with 10-25μM tubulin (Figure 5—figure supplement 2A(i)). The cooperativity of 4 αβ-tubulin dimers is predicted in the transition state (Figure 5—figure supplement 2A(ii)), in agreement with our experimental data.

To gain deeper understanding of how γ-TuRC nucleates a MT, we characterized the dynamics of αβ-tubulins from individual model simulations. When the simulation begins, we find that many αβ-tubulins interact with the open γ-TuRC longitudinally, or with an existing αβ-tubulin in a pf after its assembly onto γ-TuRC (Figure 5B, left insets). While many of these association events happen rapidly, these αβ-tubulins are only bound by the longitudinal bond energy and then dissociate from γ-TuRC or the γ-TuRC bound αβtubulin. On the other hand, αβ-tubulin dimers persist when assembled on a corner site in a MT lattice that has both longitudinal and lateral contacts. These events drive MT assembly, as has been shown by many works in the last decade (Ayaz et al., 2014; Mickolajczyk et al., 2019; Rice et al., 2019). At the sharp transition when MT nucleation occurs (Figure 5B, right insets), many αβ-tubulin dimers in the simulations stochastically assemble on neighboring sites on γ-TuRC and the transition to a closed γTuRC state becomes favorable. In other words, upon γ-TuRC closure, the favorable free energy from lateral interaction between αβ-tubulins overcomes the energy penalty from its conformational change. Combining the results from 2,119 simulations reveals that on average 5.2 ± 1 total αβ-tubulin dimers are present with γ-TuRC at this transition state (Figure 5D, left). Most strikingly, these αβ-tubulins arrange on neighboring sites in laterally-arranged groups of 2-5 αβ-tubulins (Figure 5D, right). The most probable transition state has 4 αβ-tubulin arranged on neighboring sites that form 3 lateral bonds, and this physically represents the power-law exponent measured from the average kinetics of MT nucleation.

Finally, our simulations are also able to predict the kinetics of MT nucleation from a closed γ-TuRC conformation and quantitatively predicts the measured MT nucleation dynamics from blunt seeds (Figure 5—figure supplement 2B). In summary, our computational model provides essential insights into the reaction scheme and transition state of MT nucleation from γ-TuRC.

Probing the conformation of γ-TuRC before and after microtubule nucleation

In agreement with the recent cryo-EM structures, our revised manuscript supports a model in which a conformational change in γ-TuRC from an open to a closed state occurs during the rate-limiting transition step for MT nucleation.

A high resolution cryoEM structure of the γ-TuRC-nucleated MT-end will be needed to specifically demonstrate what conformational differences in γ-TuRC occur during MT nucleation. Further capturing the transition state with a few tubulin dimers will require the development of new methods that can resolve at high resolution how tubulins assemble on γ-TuRC, and will constitute the next decades of work from the field. Despite tremendous progress in cryo-EM of the MT lattice (work by Nogales lab and others), high resolution structure of intermediates either in the MT plus-end dynamics and in MT nucleation is yet to be reached. While we are very interested in pursuing this, such an endeavor is outside the scope of this work.

Finally, we believe that the mechanism we describe is necessary to place the molecular structure of a postnucleated γ-TuRC and any transition state into a mechanistic scheme. As such, our biochemical investigation stands independently as a valuable and timely contribution to the current literature. The work presented here is a critical step toward understanding MT nucleation, which we expect the field will address in depth in the next decade.

Role of putative activators in γ-TuRC mediated nucleation

Previous methods used to identify factors that alter the function of γ-TuRC have been indirect and could not distinguish between γ-TuRC-mediated and spontaneous nucleation. These experiments also show a large amount of variability and were not well suited to determine the role of factors that have small effects on MT nucleation. E.g. published results on the effect of NME7 reported a minor, 2.5-fold, increase in activity (Liu et al., 2014), while a high variability in the role of CDK5RAP2’s g-TuNA motif has been reported: 1.7-fold in Liu et al., 2020 and 7-fold in Choi et al., 2010. Further, γ-TuRC purified with CDK5RAP2’s g-TuNA motif and NME7 still forms an open configuration (Liu et al., 2020; Wiezcorek et al., 2020; Consolati et al., 2019), raising the question how this structural data can be reconciled with the previous biochemical measurements.

Here, we used an experimental setup that allows direct visualization and measurement of γ-TuRCmediated MT nucleation, as well as the ability to distinguish these nucleation events from spontaneous ones, and assessed the role of these putative factors. We examined the role of several putative activation factors in regulating γ-TuRC’s conformation and nucleation activity. First, the γ-TuRC nucleation activation domain (g-TuNA) from Xenopus laevis protein CDK5RAP2 does not significantly increase MT nucleation activity from γ-TuRC (Figure 6A-B). NME7, another proposed activator of γ-TuRC-mediated nucleation (Figure 6B and Figure 6—figure supplement 1) did not significantly affect γ-TuRC-mediated nucleation either. Similar results were observed for TPX2 (Figure 6C-D).

Discussion of the regulation of γ-TuRC-mediated nucleation

Here we would like to address the final point raised in response to our appeal letter by the reviewers.

The current view is that there is a good reason for having γ-TuRC in an inactive conformation: to spatially restrict nucleation. Otherwise γ-TuRC would nucleate MTs anywhere in the cell, once tubulin starts binding. […] This would still not be compatible with the fact that in cells nucleation occurs from a minority of γ-TuRCs at MTOCs and not the large majority of γ-TuRCs in the cytosol.

We thank the reviewers for raising this important point. First, we note that despite their open conformation, purified γ-TuRCs nucleate MTs efficiently as shown here. Similarly, γ-TuRCs present in the cytosol in vivo also nucleate MTs efficiently. This is best exemplified by branching MT nucleation that occurs in the cytoplasm, generates thousands of MTs (Petry et al., 2013; Alfaro-Aco et al., 2017; Thawani et al., 2018), constitutes a major pathway for MT nucleation and drives spindle assembly in many organisms compared to the centrosomes (Decker et al., 2018; David et al., 2019). We propose that native, open γ-TuRC could function and be regulated in an alternative way by tuning the availability of αβ-tubulin subunits. Concentrating tubulin on MTOCs such as centrosomes (400μM tubulin, Baumgart et al., 2019),on branched MTs by TPX2 (King and Petry, 2019), or directly on γ-TuRC via XMAP215 as shown here and in a previous work (Thawani et al., 2018) could regulate MT nucleation. Cytosolic γ-TuRC’s activity could also be regulated by sequestering αβ-tubulin by Stathmin or MCAK in the cytosol and later releasing it during the cell cycle through phosphorylation. This could allow for spatiotemporal control of MT nucleation.

Second, there is little experimental evidence supporting the widely-believed hypothesis that gTuRC conformation is modified to a closed one by activation factors at the MTOCs. While g-TuSC rings on the spindle pole body of a yeast cell have a smaller diameter (Kollman et al., 2015), this was observed after the g-TuSC rings have nucleated MTs, which is consistent with the proposition we outlined above. Further, several factors proposed as putative activators of γ-TuRC have since been shown to either play a role in the assembly of γ-TuRC or to function as localization factors (Kollman et al., 2010; Leung et al., 2019; Liu et al., 2020).

Notably, the possibility of γ-TuRC closure by αβ-tubulin and its implications on MT nucleation have also been discussed in the recent cryo-EM papers (Liu et al., 2020; Consolati et al., 2019), in consensus with our model. We discuss this points in our revised manuscript in the Discussion section.

“In this model, locally concentrating soluble αβ-tubulin could upregulate the levels of γ-TuRC-mediated MT nucleation, e.g. as recently shown through accumulation of high concentration of tubulin dimers at the centrosome by MAPs55,58 and by co-condensation of tubulin on MTs by TPX2 during branching MT nucleation46, and finally via specific recruitment of tubulin on γ-TuRC through the binding of XMAP215 as shown here29,48.”

Finally, in light of several comments from the reviewers, we have drastically re-structured the manuscript and modified the Discussion section. We now first present our findings on characterization of γ-TuRCmediated, spontaneous and seed-mediated nucleation (Figure 1, Figure 2, Figure 3 and related supplements). We then discuss biochemical features of γ-TuRC that determine its nucleation activity and develop simulations that explain how γ-TuRC nucleates a MT (Figure 4, Figure 5 and related supplements). Lastly, we examine the role of several MT-associated proteins in γ-TuRC-mediated nucleation (Figure 6, Figure 7 and related supplements).

Reviewer #1:

The authors use γ-TuRCs purified from Xenopus egg extracts in combination with single molecule microtubule nucleation assays to gain insight into the process of microtubule nucleation. The key findings are that γ-TuRCs enhance microtubule nucleation by promoting the lateral association of a/β-tubulin molecules and that nucleation from γ-TuRCs is further promoted by XMAP-215. The experiments are well-executed and the data is nicely presented (although I have some questions about N numbers – see below). Overall, I think there are some interesting insights into γ-TuRC-mediated nucleation and the experiments are at the cutting edge, but I feel that we do not learn a great deal more about the role of γ-TuRC and XMAP-215 in microtubule nucleation to justify publication within a broad spectrum high-impact journal such as eLife. Several results or notions are already known e.g. γ-TuRCs and XMAP-215 promote microtubule nucleation, a higher concentration of tubulin is required for nucleation than for microtubule elongation, γ-tubulin forms filaments, XMAP-215 promotes nucleation, XMAP-215 associates with γ-TuRCs. Moreover, the main message, that γ-TuRCs promote lateral association of tubulin dimers, is widely assumed based on γ-TuRC structure and because longitudinal a/β-tubulin contacts are believed to be stronger than lateral contacts. As the authors themselves state, "the insights on MT nucleation by γ-TuRC and XMAP215 provide an essential basis to build upon". I therefore feel that, without further advances, such as how γ-TuRCs can be activated to better promote microtubule nucleation, the paper would be better suited to a more specialised journal, such as JBC.

We thank the reviewer for their insightful comments that helped us improve the manuscript significantly by performing several new experiments and interpreting our results in light of the cryo-EM structures of γtubulin ring complex. Here, we clarify several points noted by the reviewer to reflect the advance provided by our work.

First, while γ-TuRC has been widely assumed as the MT nucleator, its activity has been reported to be low, and often comparable to spontaneous MT nucleation (Kollman et al., 2010; Choi et al., 2010; Liu et al., 2014; Thawani et al., 2018). However, to directly compare the activity of γ-TuRC-mediated and spontaneous nucleation, it is necessary to assay these activities side-by-side using the same experimental setup, as well as live visualization where any spontaneously nucleated MTs in γTuRC-reaction can be distinguished from γ-TuRC-nucleated ones. Additionally, while this observation has been made for MT assembly from seeds (Wieczorek et al., 2013), it has never been demonstrated that MT nucleation from γ-TuRC required higher tubulin concentration than MT polymerization before our work. Instead, a number of reviews (Kollman et al., 2011; Roostalu and Surrey, 2017) had suggested that the dynamics of MT nucleation from γ-TuRC template could resemble the plus-end of a MT template. In our work, we developed sensitive assays and quantitative metrics to define the nucleation efficiency and rate-limiting step of γ-TuRC at the single molecule level, which for the first time could be directly compared to spontaneous and MT seed based nucleation. The results obtained and reported in our manuscript regarding this comparison are novel. Most importantly, our data on the composition of the ratelimiting, transition state consisting of γ-TuRC and four tubulin dimers. Together with the computational modeling, our works provides gives first insight into the dynamics and transition state of γ-TuRC-mediated MT nucleation, where laterally-associated tubulin dimers promote an open to closed transition in γ-TuRC conformation.

Second, as described in the common response above, we comprehensively investigated how γTuRC nucleation occurs by pairing experimental measurements and computational modelling. We characterized the biochemical features that define γ-TuRC-mediated nucleation, and find that the longitudinal γ-/αβ-tubulin interaction affinity is high. This high affinity promotes γ-TuRC’s nucleation activity by increasing the dwell time of αβ-tubulin dimers.

Third, we directly test the effect of specific MAPs on γ-TuRC-mediated nucleation. We believe that testing the activity of two previously proposed putative activation factors, CDK5RAP2’s γ-TuNA motif and NME7, with a sensitive, direct measurement in this paper is important. The finding that these factors do not significantly affect MT nucleation by γ-TuRC is significant. This result clarifies many contradicting results for the roles of these factors from previous works, where γ-TuNA was both found to affect nucleation by 7-fold (Choi et al., 2010) or minimally (Liu et al., 2020), while neither γ-TuNA nor NME7 present in complex with γ-TuRC altered the structure of the γ-TuRC from an open conformation to closed one (Liu et al., 2020; Wieczorek et al., 2020; Consolati et al., 2019). These results suggest that these MAPs do not alone function as activators, but may instead support the assembly or localization of γ-TuRC. They also highlight that MT nucleation in the cell may be regulated primarily through the availability and distribution of αβ-tubulin dimers, as is discussed in the reviewed manuscript. Only XMAP215 functions as the bona-fide co-nucleation factor with γ-TuRC and increases MT nucleation efficiency by 30-fold. These measurements had to be made side-by-side with our sensitive, direct assay to assess γ-TuRC’s nucleation activity in order to reveal the role of any of these factors in MT nucleation.

Finally, we emphasize that this information could not be obtained simply from solving EM structures, as recently achieved, as they only captured the pre-nucleation state and did not reveal anything about the nucleation reaction. All these insights could only be obtained from single molecule light microscopy paired with reaction kinetics analyses, which now constitute the state-of-the-art measurements. The transition state of γ-TuRC with four αβ-tubulin dimers represents an important milestone in understanding the γ-TuRC-mediated MT nucleation reaction scheme and its steps.

More specific comments:

1) The title should be more specific to the work done, as the current title indicates that the authors have solved the entire mechanism of nucleation.

We thank the reviewer for this comment and have modified the title of the paper to, “The transition state and regulation of microtubule nucleation γ-TuRC revealed by single molecule microscopy” to specifically reflect the work done, and additionally welcome any alternate suggestions for the title.

2) Abstract: "the underlying mechanism largely remains a mystery". This is a difficult statement to swallow. I would argue that the underlying mechanism (templating the addition of tubulin dimers into a tube-like structure) has been widely assumed (although not directly proved) for a long time, in particular after the 2010 Nature paper from David Agard's group that showed the structure of the yeast "γ-TuRC". This was good proof that the template model was correct, rather than the protofilament model. I think the authors should therefore modify their statement to something like: "While it has been assumed that…., here we provide direct evidence for….".

We thank the reviewer for this suggestion. We made major changes to the Abstract to incorporate the specific suggestion. Also, the Abstract is now more focused on the new quantitative measurements made in this work that constitute the major advance compared to the previous literature.

3) Abstract: "…we uncover that γ-TuRC nucleates a MT more efficiently than spontaneous assembly". This was already known – many studies have shown (using more standard microtubule nucleation assays) that microtubules form more readily in the presence of γ-TuRC.

While some previous works (Zheng et al., 1995; Oegema et al., 1999) reported γ-TuRC’s nucleation activity to be higher than spontaneous nucleation, a number of recent works have described γTuRC-mediated nucleation to be inefficient and similar to the levels of spontaneous nucleation in their assays (Kollman et al., 2010; Choi et al., 2010; Liu et al., 2014; Thawani et al., 2018; Liu et al., 2020). Some proportion of the nucleated MTs in all reports could be due to spontaneous assembly, which cannot be distinguished from γ-TuRC-nucleated MTs in the traditional, endpoint MT nucleation assays. In contrast, we precisely measured the nucleation activity for γ-TuRC-mediated and spontaneously nucleated MTs, which can be further distinguished by observing the growth of MT minus-end for the latter case but not the former. Such a side-by-side comparison in a sensitive assay was missing and provides insight into how γ-TuRC nucleates a MT. To highlight the most critical result from our measurements, we now state in the Abstract, “Whereas spontaneous nucleation requires assembly of 8 αβ-tubulins, nucleation from γ-TuRC occurs efficiently with a cooperativity of 4 αβ-tubulin dimers.”

4) Figure 1B: Three γ-TuRC nucleation sites are indicated by arrows – is this all of the γ-TuRCs present within this field (Can the authors show a two-colour image overlay)? If there were only three γ-TuRCs, it would not fit with the finding that spontaneous nucleation does not occur at 10.5μM tubulin….

We thank the reviewer for pointing this out and realized that this representation could be improved. All MTs nucleated in the field of view in Figure 1B were nucleated from γ-TuRC molecules. We now represent all MTs nucleated already in the first frame with yellow arrows and all new nucleation events between first and last frames with blue arrows. We additionally compared MT nucleation with and without γ-TuRC sideby-side in an experiment requested by Reviewer 3 (revised Figure 3—figure supplement 1B-C). This confirmed that γ-TuRC-mediated nucleation occurs efficiently at 10.5μM tubulin, while no MTs are nucleated spontaneously (i.e. without γ-TuRC).

5) Subsection “Defining the microtubule nucleus on γ-TuRC”: "Surprisingly, γ-TuRC nucleated MTs starting from 7 μM tubulin (Figure 1D), which is higher than the minimum tubulin concentration (C*) needed for growth at pre-formed MT plus-ends (C* = 1.4μM, Figure 1E)". I don't think this is surprising, given the results of Wieczorek et al., 2015 – these authors used solution exchange to show that, while nucleation from a microtubule seed required ~6μM tubulin, microtubules could continue to grow at ~1μM tubulin. This statement should be toned down and the authors should refer to Wieczorek et al. results.

We appreciate this point by the reviewer and have removed the word “surprisingly” from this statement in the manuscript. In our work, we directly compared γ-TuRC-mediated MT nucleation and seed-mediated assembly. Our results for seed-mediated MT nucleation differ from those of Wiezcorek et al., such that MT seeds nucleate near the critical tubulin concentration (Figure 3C-E). The differences in results from those of Wiezcorek et al., are due to different assay conditions, as detailed in the response to point (7) by the reviewer. The tubulin concentration needed for nucleation from γ-TuRC in this work cannot, therefore, be directly compared with Wiezcorek et al. results where a different assay condition produces the kinetic barrier and requirement of high tubulin concentration. For this reason, we performed the seed-mediated nucleation assay is our assay conditions to compare side-by-side with γ-TuRC-mediated nucleation in the same conditions. We request the reviewer to refer to our response for point (7) below for an in-depth explanation.

6) Figure 1G: The authors use power-law calculations to calculate that ~3.7 tubulin dimers are required for nucleation. It is not clear exactly how they did this. Given that this power analysis underlies the main finding of the paper, I think it deserves a full explanation.

We thank the reviewer for these suggestions. In the revised manuscript, we included a new subsection, “Power-law analysis of critical nucleus size on γ-TuRC” with a detailed explanation for the power-law analysis including the governing equations for the reaction kinetics balance. We further included similar analysis details for spontaneous nucleation and nucleation from MT ends in subsection “Preparation, microtubule assembly from blunt microtubule seeds and data analysis” and subsection “Size exclusion chromatography of γ-tubulin and αβ-tubulin” respectively.

7) Subsection “Does γ-TuRC nucleate a microtubule via the blunt plus-end model?”: The authors use microtubule seeds to compare their nucleation efficiency to γ-TuRCs and find that microtubules can assemble from these seeds at only 2.45μM tubulin. This is significantly lower than that found by Wieczorek et al., 2015 (6μM tubulin) – can the authors comment on this and suggest an explanation?

We thank the reviewer for this question, which echoes the related point (5). The differences between our results and those of Wiezcorek et al., 2015 are due to the different assay conditions, which we elaborate on below.

This assay was performed under different conditions by Wiezcorek et al., 2015. First, the preparation of our coverslips relies on covalently attaching PEG molecules to the glass, which is superior than the non-specific passivation of coverslips used in other works including Wiezcorek et al., 2015 as well as our previous works (Thawani et al., 2018). Second, there are specific differences in the buffer composition used. As we were initially surprised by these findings as well, we repeated this experiment exactly as published (Author response image 1A). Under the assay conditions reported in Wiezcorek et al., we observed similar results where most MT seeds do not assemble polymerizing MT at 5μM tubulin, while at 7μM tubulin, MTs assemble from seeds (Author response image 1A). This suggests that different assay condition or coverslip preparation results in distinct nucleation kinetics from seeds in vitro. Further, we characterized which assay condition represents that the barrier in assembly from seeds exists in a physiological context. We examined how MT assembly from seeds occurs in Xenopus egg extracts. Using the same coverslip preparation and MT seed attachment as Wiezcorek et al., we added Xenopus egg extracts instead of purified tubulin to visualize MT assembly. Surprisingly, MTs assembled immediately from the seeds (Author response image 1B). We note that MT nucleation by seeds in extract is in contrast with γ-TuRC-mediated nucleation in egg extract, where a large number of γ-TuRCs are present in the cytoplasm, yet MT nucleation from these molecules is not observed without the addition of MTOCs or branching effector RanGTP (Petry et al., 2013; Thawani et al., 2018; Alfaro-Aco et al., 2017). In sum, the nucleation barrier for a blunt-to-tapered transition does not appear as the major contributor to the kinetics of γ-TuRC-mediated assembly either our in vitro setup or in Xenopus egg extracts.

Author response image 1

Therefore, when repeated side-by-side with the same assay conditions, MT seeds do not display a nucleation barrier while γ-TuRC does. To clarify this point in the manuscript, in subsection “Contribution of end architecture of γ-TuRC to microtubule nucleation”, we state, “Notably, the distinct minimal tubulin concentration needed for seeds to assemble a MT as compared to a previous report24 results from the differences in assay conditions.”

8) Subsection “Molecular insight into microtubule nucleation by γ-TuRC”: Experiments to nucleate microtubules from γ-tubulin oligomers: I don't think this adds much to the paper. It is already known that γ-tubulin can form filaments and it is hard to understand how these filaments would nucleate microtubules, given that the filaments are presumably made of laterally contacting γ-tubulin molecules, thus creating a linear, rather than circular, template for the addition of a/β-tubulin dimers. It was also not clear from the methods what the final salt concentration was during the nucleation reactions, given that the g-tubulin was diluted in high salt buffer before being added to the reaction buffer. This is important as the authors show that tubulin oligomers do not form at high salt concentrations. If the authors want to keep these experiments in the paper, they need to cite work from the groups of Alvarado-Kristensson, Binarova and Monasterio who have shown that γ-tubulin can oligomerise into filaments.

We now cited the works suggested by the reviewer in subsection “γ-tubulin has a high affinity for αβ-tubulin” and specified the final salt concentration in the assay in subsection “Nucleation of microtubules from purified γ-tubulin”, “varying concentration of γ-tubulin was prepared by diluting purified γ-tubulin in a high salt buffer (50mM K-MES pH 6.6, 500mM KCl, 5mM MgCl2, 1mM EGTA), […] to a final salt concentration of 44mM KCl”.

With further analyses of our negative stain EM data (Figure 4—figure supplement 2C-E), we show that γ-tubulins within the filament arrange into laterally-associated arrays. We speculate that MT generation could occur from assembly of a αβ-tubulin dimers onto the γ-tubulin arrays, followed by further polymerization and closure of the tube to form a MT. Notably similar mechanisms involving a sheet-to-tube transition via a sheet-closure have been previously proposed for MT assembly (Wu et al., 2009; Vitre et al., 2008; Voter and Erickson, 1984). Finally, we agree with the reviewer that this is not a major point in the manuscript and have therefore, moved these data to the supplementary figures.

9) Subsection “Molecular insight into microtubule nucleation by γ-TuRC”: The stochastic models, used to predict how many γ-tubulin sites must to be occupied to have the best chance of 2 adjacent tubulin dimers being present, are based on a perfectly circular arrangement of γ-tubulin molecules (Figure 2H). Liu et al., 2019 have just shown the cryo-EM structure of the Xenopus γ-TuRC. It is in an open confirmation, where the first and last γ-tubulin molecules are not adjacent to each other. I presume this would change (albeit perhaps only slightly) the calculations from the stochastic model simulations, so the authors should repeat them using an open ring where there is no lateral contact if positions 1 and 13 are occupied.

We thank the reviewer for this comment. In the revised version, we developed Monte Carlo simulations to model the process of MT nucleation from γ-TuRC comprehensively. We describe the simulation methods and results in detail in the common response. Briefly, we incorporated γ-TuRC’s open conformation, the precise geometry of the seam, as well as stochastic association and dissociation of αβ-tubulin dimers to γTuRC’s sites or to other αβ-tubulins. This geometry, encoded in our detailed simulations, incorporates the reviewer’s suggestions. This model independently recapitulates the detailed dynamics of γ-TuRC-mediated MT nucleation of our measurements, and also identifies the arrangement of αβ-tubulin dimers in the transition state as described in the common response. We hope that the new results from the detailed simulation model addresses the points raised by the reviewer.

10) For quantifying the numbers of microtubules nucleated in e.g. Figure 1F, Figure 3A,3C it is not clear how many times the experiments were performed and whether the graphs represent mean or median values. Given the shakiness of the lines in the graph in Figure 3A, the N numbers seem low. I would expect that the experiments (being quick to perform) would have been performed several times (perhaps 5 to 10 times) and that a mean or median value was used to represent microtubule numbers at each timepoint. Can the authors confirm this, or do more repeats?

We appreciate this point raised by the reviewer and addressed it in the following way:

First, we performed and included additional experimental repeats and analyses. Between 3-5 repeats were performed for most MT nucleation experiments. We expanded on our description to also provide detailed information on the number of repeats performed in every figure legend. Notably our power-law analyses with 3-5 pooled replicates result in an estimate of 3.9 ± 0.5 tubulin dimers for γ-TuRCmediated nucleation, 8.1 ± 0.9 for spontaneous nucleation, 1 ± 0.3 for assembly from blunt MT end, and 3.3 ± 0.8 for XMAP215/γ-TuRC-mediated co-nucleation.

Second, each replicate for an experiment was performed with independent γ-TuRC purifications where the concentration of γ-TuRCs obtained varies between purifications. The variation in the absolute number of MTs nucleated between replicates mostly represents variance in number of γ-TuRCs. Therefore, reporting the mean or median value was not appropriate. In light of the reviewer’s comment, we instead developed new ways of combining data from independent replicates using the following two approaches. (i) For data where power-law analysis of nucleation versus concentration was performed (Figure 2C-D, Figure 3B, Figure 3D-E, Figure 7D-E), we reported a representative curve the nucleation kinetics to display the raw data and combined data from all repeats in the power-law analysis (Figure 2D, 2B, 2E, Figure 7E). (ii) To combine data in the power-law analysis from multiple γ-TuRC experiments, it was necessary to set a normalization factor that we specify in individual figure legend. For example, in the legend for Figure 2B, we state, “The rate of nucleation at 10.5μM was set to 1 to normalize differences in γ-TuRC concentration from individual experiments.” For experiments to study the effect of MAPs (Figure 6B, 6D, Figure 7B, 7F), we combined results from multiple experiments, as requested by the reviewer, by setting a normalization factor between different experimental repeats and displaying kinetics from all repeats on the plot. For example, in the legend for Figure 6D, we state, “To account for the variable γ-TuRC concentration across purifications, the number of MTs nucleated in control reactions at 150 seconds was set to 1. All data was pooled and reported.” We also report 95% confidence intervals on our data as detailed in Methods and individual figure legends.

Finally, we emphasize that these are the most challenging and time-intensive experiments we have witnessed in our careers. Getting these to work was a breakthrough in itself. We observed that some γTuRC’s dissociate and lose activity during freeze-thaw cycle, and therefore, we performed each assay with freshly purified γ-TuRC. Our most important metric was reproducibility, which was ensured with all experiments reported and we specify this in detail for individual figure panels.

11) Figure 3A: the authors show that TPX2 has no role in microtubule nucleation from γ-TuRCs, but previous studies (Wieczorek et al., 2015, Roostalu et al., 2015, Woodruff et al., 2017) have shown that TPX2 does promote nucleation. Do the authors believe there is a difference in the requirement of TPX2 when considering γ-TuRCs versus microtubule seeds as templates? Can the authors repeat previous results showing that TPX2 promotes nucleation from microtubule seeds, as a positive control to show that their purified TPX2 is functional?

We thank the reviewer for raising this important point and address it as follows. First, we tested the effect of TPX2 on γ-TuRC’s nucleation activity at a higher TPX2 concentrations (30-50nM; Author response image 1C–D). While the TPX2 appeared to generate more MTs than γ-TuRC alone (Author response image 1C), the formation of large tubulin clusters occurred at these higher TPX2 concentration and MTs were seen to be generated from these clusters (Author response image 1D). The finding that TPX2 can co-condense with tubulin and generate MTs spontaneously is in agreement with previous work (King and Petry, Nat Comm 2020). Such a high concentrations of TPX2 (~50nM), above the co-condensation boundary, was also used in previous works where TPX2 was observed to nucleate MTs spontaneously when high TPX2 concentrations were used (Roostalu et al., 2015, Woodruff et al., 2017). However, a lower concentration of TPX2 (~1nM) is sufficient to bind to the MT lattice and does not form tubulin clusters (Roostalu et al., 2015; King and Petry, 2020). Because we wanted to measure the effect of soluble TPX2 on γ-TuRC’s nucleation activity and to avoid confounding measurements via tubulin clustering, we operated below this phase boundary at 10-20nM TPX2. At this concentration, TPX2 completely coated the MT lattice, yet did not have a significant effect on γ-TuRC-mediated nucleation (Figure 6C-D).

In summary, TPX2 does not significantly affect γ-TuRC-mediated nucleation, whereas its high concentrations are required for spontaneous MT assembly. Several studies from our lab have previously shown that in Xenopus egg extracts, TPX2 also does not directly increase MT nucleation from γ-TuRC. This is the case at TPX2’s physiological concentration (30nM) as well as at high concentration (1μM) (Alfaro-Aco et al., 2017; Thawani et al., 2019).

12) Subsection “How do γ-TuRC and XMAP215 synergistically nucleate microtubules?”: "XMAP215 effectively decreases the minimal tubulin concentration necessary for MT nucleation from γ-TuRC to 1.6 μM". How was this calculated? Also, does adding XMAP215 also increase the probability of spontaneous nucleation? In a model where XMAP215 binds γ-tubulin and uses TOG domains to add tubulin dimers for the initial nucleation event, the addition of XMAP215 should not increase the probability of spontaneous nucleation.

We appreciate these two questions. First, in Figure 7—figure supplement 1C, we display a set of experiment where constant γ-TuRC and XMAP215 concentration was used, while tubulin concentration was varied to allow a measurement of the critical nucleus size. XMAP215 and γ-TuRC together nucleate MTs starting from a tubulin concentration of 1.6μM, as quantified in Figure 7D. To address the second question, we compared side-by-side how many MTs are nucleated by XMAP215 alone, γ-TuRC alone and γ-TuRCs with XMAP215 (Figure 7—figure supplement 1A), which were quantified in Figure 7—figure supplement 1B. We find that at 7μM tubulin, neither γ-TuRC nor XMAP215 alone significantly nucleate MTs, yet robust MT nucleation only occurs when XMAP215 and γ-TuRC are present together. We hope this addresses the reviewer’s question.

13) Figure S4B: the authors test whether the C-term γ-tubulin-interacting region of XMAP215 promotes microtubule nucleation, and show that it does not. It would be informative to also test whether the rest of XMAP-215 alone (i.e. XMAP-215 minus the γ-tubulin binding region) can promote microtubule nucleation from a γ-TuRC. Presumably not, but this truncated form of XMAP-215 should in theory be able to bind to microtubule plus ends via the TOG domains. It would therefore be a nice experiment to show that it is not plus-end bound XMAP-215 that is promoting nucleation, but it is the XMAP-215 that binds to the γ-TuRC.

XMAP215 construct containing TOG domains 1-4, and further repeats for C-terminal truncation TOG5CT. The results are displayed in Figure 7—figure supplement 1D-E and we compared these side-by-side with the activity of full-length XMAP215 (positive control) and γ-TuRC alone (negative control). To summarize, we find that neither the C-terminal, which binds γ-tubulin directly (Thawani et al., 2018), nor the Nterminal truncations of XMAP215 increase γ-TuRC-mediated nucleation significantly, while wild-type XMAP215 drastically induces MT nucleation from γ-TuRC in vitro.

Reviewer #2:

The present manuscript uses purified γ-TuRC, tubulin, and different MAPs, to reconstitute microtubule nucleation in vitro, and to quantify nucleation rates, growth speed, and cooperativity of tubulin dimers. Using coated coverslips to anchor tagged γ-TuRCs, in combination with TIRF microscopy, several important observations are made:

- Nucleation of microtubules occurs from γ-TuRCs de novo, and this is more efficient than spontaneous assembly.

- The critical nucleus for nucleation from γ-TuRCs comprises only 3-4 tubulin dimers, which is less than half the size required for spontaneous assembly.

- γ-TuRC-mediated nucleation differs from plus-end growth at blunt microtubule ends, where a single tubulin dimer suffices as a nucleus.

- γ-TuRC-mediated nucleation is not influenced by TPX2. By contrast, XMAP215 decreased the minimal tubulin concentration and increased the nucleation rate, by binding first to the γ-TuRC, yet requiring a similar critical nucleus of 3-4 tubulin dimers.

Altogether, these data provide extremely valuable insight into the mechanisms of microtubule nucleation, since they clarify multiple major issues: γ-TuRCs act directly as templates for microtubule nucleation, ruling out other hypotheses that propose spontaneous nucleation, followed by capping of the free minus-ends by γ-TuRCs. Most importantly, we have for the first time a concrete idea about the critical size of the tubulin nucleus, and about the involvement (or non-involvement) of XMAP215 and TPX2 in the nucleation process.

The manuscript is timely, since several other studies on the γ-TuRC (e.g. structure of the native full complex) are likely to be published by other groups in the very near future.

We thank the reviewer for recognizing the breakthrough and timeliness of our work.

Reviewer #3:

The manuscript by Thawani et al., together with a recently posted study by the Surrey group on the bioRxiv preprint server, is the first to study microtubule nucleation from purified, immobilized gTuRC at the single molecule level by TIRF microscopy. Using this approach the authors investigate and compare nucleation from gTuRC with spontaneous MT assembly and nucleation from preformed MT seeds. They find that nucleation from gTuRC is more efficient than spontaneous assembly, but less efficient than nucleation from MT seeds. They argue that this is due to relatively weak longitudinal affinity between g-tubulin and ab-tubulin based on measurements of ab-tubulin interaction with purified g-tubulin and ab-tubulin in biolayer interferometry assays. XMAP215 significantly increases gTuRC nucleation activity. They estimate that nucleation from gTuRC alone requires 3-4 ab-tubulin dimers to bind to gTuRC before the kinetic nucleation barrier is overcome and argue that this is facilitated by the lateral array of g-tubulins in the gTuRC.

The establishment of TIRF imaging of nucleation from gTuRC at single molecule level is overdue and the presented work is a great achievement in this regard, providing interesting new ideas and experimental possibilities. However, the authors base their conclusions on various assumptions that cannot be made without further evidence and control experiments. A major issue is the fact that native gTuRC is not a perfect template that matches the symmetry of a microtubule, as revealed by three independent recent Cryo-EM studies by the Kapoor, Schiebel and Surrey labs. Overall, the study requires significant additional work that in my opinion would go beyond what would be reasonable for a revision in eLife.

We thank the reviewer for the positive comments recognizing the breakthrough of this work and for providing critical feedback of how this work can be advanced further to provide deeper mechanistic insight and reconcile it with the recent cryo-EM structures. As described in the common response above, we reevaluated the previous assumptions, added several entirely new experiments and developed computational models to provide deeper insight how a MT is nucleated by γ-TuRC based on the recent cryo-EM structures. In the following, we directly address all comments raised by the reviewer.

Major points:

1) Subsection “Reconstituting and visualizing microtubule nucleation from γ-TuRC”: "MT nucleation events occurred specifically from single gTuRC molecules" – gTuRCs are not labelled in this assay, how do the authors get to this conclusion? How can they exclude that some of the MTs are from spontaneous nucleation?

We thank the reviewer for raising this crucial point, which we address in the following response. First, we included new data in which purified γ-TuRCs were covalently attached to both biotin- and Alexa-568 fluorescent-dye (Figure 1C and Figure 1—figure supplement 1C). MT nucleation events were always observed from individual Alexa-568 labelled, green spots (Figure 1C) demonstrating that individual γ-TuRC molecules nucleate MTs and significant spontaneous nucleation does not occur. We note that many experiments in this work were not performed with fluorescent γ-TuRC because we empirically found that additional labelling of γ-TuRC with the dye using a cysteine reaction resulted in fewer γ-TuRCs and lower activity at the end of the purification, vis-a-vis from biotinylated γ-TuRCs only (data not shown). Therefore, we used biotinylated γ-TuRCs for most experiments which was sufficient to answer our questions. Second, and of primary importance, we note that the characteristics of MTs nucleated by γ-TuRC and those nucleated spontaneously can be clearly distinguished using our live assay. In Author response 4A, we show kymographs for γ-TuRC-nucleated MTs from Figure 1C and for spontaneously nucleated from Figure 3—figure supplement 1A side-by-side. We marked the minus- and plus-end of MTs with dashed and solid yellow lines, respectively to clearly demonstrate this. For γ-TuRC mediated nucleation, MTs emerge from zero length from the coverslips and pivot around the nucleation site due to the attachment. Most importantly, the minus-end of these MTs, marked by dashed yellow lines in the kymographs, does not grow while only the plus-end grows, as shown in solid yellow lines. In contrast, for spontaneously nucleated MTs, nucleation occurs from solution and significant translational and rotational motion of MTs is observed due to the lack of attachment.

Finally, both ends of MTs, minus- and the plus-ends shown with dashed and solid lines respectively, elongate. Another way to observe whether the minus-end grows or is capped is by following fiducial marks in the MT lattice that occur stochastically from to fewer fluorescent tubulin molecules incorporating into the lattice than average. In Author response image 2A, we displayed some of these fiduciary marks with red dotted lines. For MTs nucleated from γ-TuRCs, the distance from these marks to the minus-end of the MT does not change over time, while that from the plus-end increases. In contrast, for spontaneously nucleated MTs, the distance of these fiduciary marks from both minus- and plus-end increases over time. This again shows that minus-end does not grow for γ-TuRC-nucleated MTs, while it does for spontaneously nucleated MTs. This was used to distinguish γ-TuRC-mediated and spontaneous nucleation events.

Author response image 2

Finally, as requested by the reviewer in point (6) below, we compared γ-TuRC-mediated and spontaneous nucleation side-by-side (Figure 3—figure supplement 1B-C). As described in the manuscript, few spontaneous nucleation events occur in comparison to γ-TuRC mediated nucleation events with the tubulin concentrations used here. We elaborate on this further in our response to point (6).

2) The main conclusions of this study regarding the nucleation mechanism are based on the assumption that gTuRC is a perfect template, which has recently been disproven. Cryo-EM data of purified native human and Xenopus gTuRC clearly show that the g-tubulin array in gTuRC does not match the symmetry of a MT, by displaying lateral gaps between g-tubulin molecules, in particular in the part of gTuRC that contains GCP4, 5, and 6. This would be expected to impact its ability to promote lateral interactions between ab-tubulin dimers.

We thank the reviewer for raising this important point. In the revised manuscript, we consider the implications of the γ-TuRC’s conformation revealed by the cryo-EM structures, as described in the common response above. Altogether, our revised work shows that γ-TuRC’s nucleation properties are defined by the thermodynamic barrier from the open conformation of native γ-TuRC which does not allow complete lateral αβ-tubulin interactions.

3) Figure 2C-E: It is assumed that γ-tubulin alone, through lateral interaction, forms a filament-like microtubule nucleation template. There are two issues here. First, there is no evidence that the g-tubulin filaments are indeed formed by lateral g-tubulin interactions. In fact, the EM image in S2C shows a fiber much thicker than expected.

We thank the reviewer for raising this point and would like to clarify that arrangement of γ-tubulin into filaments. To characterize the arrangement of γ-tubulins within filaments, we optimized the negative staining of γ-tubulin filaments to minimize background, acquired 370 micrographs and obtained 3D reconstructions of γ-tubulin filaments in RELION (Figure 4—figure supplement 2C-E), as described in the figure legend and subsection “Negative stain electron microscopy of γ-tubulin filaments”. Reconstructions revealed that the filaments are made of linear arrays of laterally-interacting γ-tubulins with a constant subunit repeat of 54Å in each array. This arrangement of γ-tubulins was also observed in the crystal contacts across unit cells in the structure of γtubulin (Aldaz et al., 2005). This subunit repeat distance does not match that between α- and βtubulins interacting longitudinally in a protofilament (40Å). In sum, γ-tubulins arrange laterally along the length of γ-tubulin filaments, an arrangement that differs from γ-tubulin arranged helically along the cross-section of a MT. Here we acknowledge personal communication with Drs. Michelle Moritz and David Agard, who observed a similar arrangement of γ-tubulin filaments and MT nucleation from these assemblies using a light scattering assay (King et al., bioRxiv 2020).

Second, the authors should present evidence that the formed ab-tubulin polymers are actually microtubules rather than other types of filaments (e.g. protofilaments or polymerization along g-tubulin filaments).

We thank the reviewer for raising this important point. We have several lines of evidence that show that the αβ-tubulin forms MTs here. First, we measured the αβ-tubulin intensity along the length of polymers generated by γ-tubulin and of MTs nucleated by γ-TuRC. We find that the mean αβ-tubulin intensity along their lengths to be similar (Author response image 3A) and polymers grow with similar plus-end growth speed (Author response image 3A). Second, we visualized the nucleation reactions described in Figure 4—figure supplement 2A with negative stain electron microscopy (Author response image 3B). While a lot of protein background was observed on the EM grids likely due to high concentration of unpolymerized αβ-tubulin, MTs were clearly seen on the grids. Third, we find that γ-tubulin oligomers attached to the coverslips are also sufficient to nucleate MTs from the coverslips (Author response image 3C), and large filaments are not necessary for this activity. Finally, as these polymers are not curved and short (less than 100nm) as previous observed for protofilaments (Portran et al., 2017), but are indeed MTs formed from αβ-tubulin.

Author response image 3

In summary, our data shows that MTs nucleate from high concentration of γ-tubulins where γtubulin assembles into filaments. While some of the data provided in Author response image 3A will be of general interest to the field, it deviates from the main focus of the current manuscript, and we will communicate this in an independent study. We believe that revealing the arrangement of γ-tubulins in the filament (Figure 4—figure supplement 2C-E) clarifies that γ-tubulin filaments and oligomers do not position MT-like plus-ends, but instead form lateral arrays that are sufficient to nucleated MTs from αβ-tubulin dimers.

4) Fig2D: Are these really protofilaments? They seem much thicker. Also, can the authors exclude that g-tubulin polymerizes during coating of the chip?

We thank the reviewer for this comment, which prompted us to comprehensively characterize γ-/αβ-tubulin interaction using a number of biochemical approaches. As described in the common response above, using size exclusion chromatography (SEC) and single molecule binding assays (Figure 4A-C), we find that the affinity between γ-tubulin and αβ-tubulin is higher than that between two αβ-tubulins. Our interferometry experiments performed with the new orientation where αβ-tubulin is bound to the probe and γ-tubulin is in solution corroborates this result (Figure 4—figure supplement 1B(ii)). As outlined in the common response, the reverse orientation used before likely resulted in an occlusion of αβ-tubulin binding site on γ-tubulin and no binding could be observed. Further, as the reviewer correctly points out, γ-tubulin associates non-specifically with the probe to a lower extent in the interferometry experiments. Thus, we think that SEC and single molecule binding assays supersede this experiment. Hence, we have moved the interferometry experiments to the supplement as supporting observations. With this revised picture, it was not necessary to display the protofilaments that form from αβ/αβ-tubulin as previous works clearly show that more favorable longitudinal αβ/αβ-tubulin interaction results in assembly of tubulin protofilaments (Portran et al., 2017).

5) Figure 2H: First, the simulation (i) assumes that gTuRC is perfectly ring-shaped (but it is not, see point 2 above) and ii) does not take into account binding affinity with tubulin dimers already bound to the template. After binding of the first dimer, binding of a second dimer at the neighbouring position may be promoted by lateral affinity. Also, I don't understand how the probability in calculated – should it not increase further after 4 sites are occupied? At this point it should be much more likely to have lateral contacts than not.

We thank the reviewer for this question. In the common response above, we outline the assumptions and results from the explicit Monte Carlo simulation model for MT nucleation from γ-TuRC (Figure 5 and related supplements). Our revised model incorporates all of the suggestions made by the reviewer. First, we assume an open conformation where lateral αβ-tubulin contacts are not promoted, as shown by the recent structural work. Second, in our stochastic simulations, we allow all possibilities of αβ-tubulin associations including those with γ-TuRC as well other assembled tubulin dimers at various sites. Third, tubulin dimers that arrive have either no lateral neighbors, half a neighbor at the seam, one or two lateral neighbors. The number of neighbors that contribute to the total lateral bond energy affect the dissociation rate of each dimer, as modeled previously for MT polymerization (VanBuren et al., 2002) and shown experimentally (Mickolajczyk et al., 2019) Finally, in Figure 5D, we characterize the arrangement of αβ-tubulin dimers at the transition state for each simulation where γ-TuRC nucleation occurs. On average 5.2±1 total αβtubulin dimers were present (n=2119 simulations). Here, many arrangements of αβ-tubulin dimers on γTuRC result in the same the total number of lateral αβ-tubulin contacts formed at the transition state, which is the important feature of our model (Figure 5D, right). Indeed, a few simulations showed a second αβ-tubulin dimer bound to one αβ-tubulin that was already assembled on γ-TuRC at the transition state. While many αβ-tubulin arrangements can occur in the transition state, the ones that exist most frequently have one layer of αβ-tubulins on γ-TuRC and are diagrammed in Figure 5D, right.

Notably these comprehensive simulations go beyond the previous model where we simply counted the number of sites occupied, and αβ-tubulin’s dissociation rate or local interactions from the neighbors were not incorporated in detail. In sum, our Monte Carlo simulations address the reviewer’s comments and comprehensively describe the dynamics of MT nucleation from γ-TuRC in detail.

6) Considering the technical challenges with experimental repetitions and use of different, freshly prepared gTuRC preps for each repetition, it would be useful to repeat the nucleation assay to measure spontaneous and nucleation from seeds and from gTuRC side-by-side to properly compare the different conditions.

We thank the reviewer for this comment and address it as follows. First, as suggested by the reviewer, we performed side-by-side comparison of γ-TuRC and spontaneous MT nucleation in Figure 3—figure supplement 1BC at 10.5μM αβ-tubulin. In Figure 3—figure supplement 1C, we display results from one repetition and supply the other one in Extra Figure 4b and in the Source data file. As described in the manuscript, γ-TuRC-mediated nucleation occurs efficiently at 10.5μM αβ-tubulin, but spontaneous MT nucleation does not. Because each experiment shown here is challenging and time intensive, it is not possible to side-by-side repeat all concentrations reported for γ-TuRC and spontaneous nucleation.

Second, due to the very nature of these in vitro experiments, they are highly reproducible from one experimental replicate to the next. In this work, we used the same batch of αβ-tubulin for comparison across experiments, buffer reagents prepared identically and state-of-the-art coverslip preparation which ensured high reproducibility for spontaneous nucleation and assembly from seeds. In particular for spontaneous MT nucleation, we have displayed the absolute number of spontaneous MTs nucleated in three independent repetitions in Figure 3B, inset. The number of MTs nucleated spontaneously with specified αβ-tubulin concentration is very similar from one replicate to the next. Similarly, reproducibility was observed acorss all three repetitions of MT assembly from seeds, where 2.45μM tubulin was sufficient for seeds to assemble MTs in each replicate. Further, the absolute rate of MT assembly was similar from one replicate to the next. Thus, in Figure 3E, we have displayed data from all three replicates without any normalization.

Notably, for γ-TuRC-mediated nucleation experiments, the variable concentration of purified γTuRC introduces some variability between individual repeats. Thus, for each replicate, that the control experiment with γ-TuRC alone at a specific concentration results in 2-4-fold variation in the number of MTs nucleation. Yet, the trends reported in the manuscript do not change from one-replicate to the next. To exemplify this, in Author response image 2C, we present all three replicates for the data corresponding to Figure 2AC. Because the concentration of purified γ-TuRCs varies roughly 2-4-fold between replicates (data sets), the number of MTs nucleated (y-axis) between individual repeats varies but data within individual experimental set can be compared where we used the same γ-TuRC purification. Despite this variation from one replicate to the next, the trends reported in the manuscript remain invariant across replicates. Minimal nucleation was seen at 3.5μM and 7μM tubulin respectively and increasing level of nucleation observed from 10.5-21μM. Therefore, to pool data from multiple replicates, we set a normalization scale (Figure 2D), which we describe in the Materials and methods section and figure legends. E.g. in subsection “Power-law analysis of critical nucleus size on γ-TuRC”, we state, “As the total number of γ-TuRC molecules obtained from different days purifications changes, the rate of nucleation from γ-TuRCs at 10.5μM tubulin was set to 1 (normalization factor) to allow pooling of all datasets for γ-TuRC-mediated nucleation.”

Finally, the power-law analyses used here is insensitive to the γ-TuRC concentration as it measures the fold-change in MT nucleation for each unit change in tubulin concentration. Hence, this measurement was used to compare across different experiments with γ-TuRC, seeds and spontaneous nucleation.

It should be discussed that the minimal concentration for nucleation from seeds in a previous study by Wieczorek et al., (2015) was quite different and similar to nucleation from gTuRC in the present study.

We appreciate this comment from the reviewer that echoes reviewer 1’s comment. Note that there are specific differences between the assay conditions used by Wiezcorek et al., 2015 and this work including different preparation of coverslips and assay conditions. Our coverslip preparation relies on covalently attaching PEG molecules to the glass, which is superior than the non-specific passivation of coverslips used in other works including Wiezcorek et al., 2015 as well as by us previously to assay MT growth (Thawani et al., 2018). As we were initially surprised by the differences of our findings compared to published work, we repeated these experiments with the same coverslip preparation and assay condition as described by Wiezcorek et al. (Author response image 1A). The resulting assay yields similar results as Wiezcorek et al. where most MT seeds do not assemble polymerizing MT at 5μM tubulin, but at 7μM tubulin, MTs assemble from seeds (Author response image 1A). In sum, with our in vitro assay conditions where we compare nucleation from seeds and from γ-TuRC side-by-side, all seeds assemble MTs at 5μM tubulin. The differences with the published work stem from the specific assay condition.

To examine which of these two in vitro conditions better represents the endogenous conditions, we subjected the blunt seeds to Xenopus egg extracts instead of purified tubulin by adapting the assay preparation from Wiezcorek et al. We visualized MT assembly from seeds by spiking fluorescent tubulin in egg extracts. Surprisingly, MTs assembled from seeds immediately. We note that this result is in contrast with γ-TuRC-mediated nucleation in Xenopus egg extracts, where that a large number of γ-TuRCs are present in the cytoplasm, yet MT nucleation from these molecules is not observed without MTOCs or branching effector RanGTP (Petry et al., 2013; Thawani et al., 2018; Alfaro-Aco et al., 2017). This comparison corroborates our in vitro results and assay conditions.

Finally, as we aimed to compare their kinetics with γ-TuRC-mediated nucleation side-by-side in our work, we used the same assay condition and coverslip preparation that we used to visualize nucleation from single γ-TuRC molecules. Under these conditions, MT seeds require low tubulin concentration to nucleate. To clarify this in the manuscript, in subsection “Contribution of end architecture of γ-TuRC to microtubule nucleation”, we state, “Notably, the distinct minimal tubulin concentration needed for seeds to assemble a MT as compared to a previous report24 results from the differences in assay conditions.”

7) It is not very clear how many times experiments were repeated and what the exact outcomes of each were. Sometimes data from different repetitions was "pooled", sometimes only one "representative dataset" of several repetitions is shown. Would it not be more appropriate to plot mean values from the multiple repetitions? The descriptions in all three figure legends are quite vague, for example Figure 2: "…were repeated at least twice with multiple supporting results…" – what exactly does this mean?

The same is true for the transparent reporting form:.…"confirmation and supporting experiments were performed with slightly different conditions and complete agreement was found amongst all measurements. The datasets were sufficient to give confidence on the measured value of interest…". What are the slightly different conditions? How are "complete agreement" and "confidence" determined?

Source data is not provided because "all data generated in this study was pooled (no data was left out)", but if only one "representative dataset" is shown, my understanding is that some data was in fact left out.

We appreciate this point raised by the reviewer and have done the following to thoroughly address it. First, performed and included additional experimental repeats and analyses as requested by reviewer 1. Between 3-5 repeats were performed for most MT nucleation experiments. We have also now provided detailed information on the number of replicated performed for every experiment provided in individual figure legend. Notably with this additional data pooled, our power-law analyses results in cooperativity of 3.9 ± 0.5 tubulin dimers for γ-TuRC-mediated nucleation, 8.1 ± 0.9 for spontaneous nucleation, 1 ± 0.3 for assembly from blunt MT end, and 3.3 ± 0.8 for XMAP215/γ-TuRC-mediated co-nucleation.

Second, as described in the response to point (6) above, each replicate was performed with an independent γ-TuRC purification. As a result, the variation in the absolute number of MTs nucleated between replicates mostly represents variation in concentration of γ-TuRCs used. Thus, reporting the mean or median value did not appropriately represent the experimental data. Instead, to present data from all the replicates as suggested by the reviewer, we developed alternative ways of combining data using the following two approaches. For data where power-law analysis of nucleation versus concentration was performed (Figure 2C-D, Figure 3B, Figure 3D-E, Figure 7D-E), we reported a representative curve the nucleation kinetics to display the raw data and combined data from all repeats in the power-law analysis (Figure 2D, Figure 3B, 3E, Figure 7E). This was reported in the figure legend precisely and raw time-series data from the other replicates not displayed in Figure 2C, Figure 3D, Figure 7D is provided in the Source data file. While replicates from seed-templated and spontaneous nucleation can simply be combined as is, to combine data in the power-law analysis from multiple γ-TuRC experiments, it was necessary to set a normalization factor that accounts for variance in γ-TuRC concentration. We specify this normalization factor in individual figure legend. For example, in the legend for Figure 2B, we state, “The rate of nucleation at 10.5μM was set to 1 to normalize differences in γ-TuRC concentration from individual experiments.” For experiments where we study the effect of MAPs (Figure 6B, 6D, Figure 7B, 7F), we combined results from multiple experiments, as requested by the reviewer, by setting a normalization factor between different experimental repeats and displaying kinetics from all repeats on the plot. For example, in the legend for Figure 6D, we state, “Number of MTs nucleated in control reactions at 150 seconds was set to 1 to account for variable γ-TuRC concentration across purifications, all data was pooled and reported.”

Third, as we performed and included further replicates, it was not necessary to refer to supporting data in most places. In few cases where we felt the need to mention the supportive data in the revision manuscript, we specified the exact differences for those additional experiments. The differences were in concentration points used. To exemplify this for Figure 4B-C’s legend, we state, “(B) Single molecule microscopy was performed with γ-tubulin and αβ-tubulin. Control buffer (left panels, (i) and (ii)) or biotinylated αβ-tubulin (right panels, (i) and (ii)) was attached to coverslips, incubated with fluorescent αβ-tubulin (i) or γ-tubulin (ii) molecules, set as 0 seconds, and their binding at 60-90 seconds. Number of bound molecules were analyzed for the first 15 seconds of observation described in Methods. Experiments and analyses in (B-C) were repeated identically two times, pooled and reported. n=56 data points each were displayed as mean ± std in the bar graph in (C). Further confirmed with a third supporting experimental set where the observation began later at 180 seconds and was therefore, not pooled.”

Fourth, we also report 95% confidence intervals on the number of MTs nucleated to quantitatively represent the error in our measurements. This is displayed with shaded regions for each measured curve. The confidence interval was calculated to represents the heterogeneity between various fields-of-view in a reaction and is detailed in Methods and individual figure legends.

Finally, we have provided source data for Figure 2B, 2C, 2D, Figure 3—figure supplement 1C, Figure 3—figure supplement 1D, Figure 3B, 3D, 3E, Figure 4C, Figure 6B, 6D, Figure 7B, 7D, 7E, 7F, Figure 7—figure supplement 1B, Figure 7—figure supplement 1E. We have also provided detailed information on sample size in the transparency reporting form, and thoroughly inspected for any other instance where information was left out.

8) What is the percentage of gTuRCs that nucleate microtubules in the TIRF assay? This number should be provided.

To address this question, we used our labelled γ-TuRC assays and either directly measured the number of MTs that nucleated from labelled γ-TuRCs in the presence of highest tubulin we tested at 21μM tubulin and additional 100nM XMAP215, or directly measured the number γ-TuRCs that nucleated with lower 10.5μM and further back-calculated the nucleation efficiency at the highest tubulin concentration tested (21μM) using previously measured curves. We find that approximately 10-15% of γ-TuRCs nucleate MTs. Notably this number is likely to be an underestimate as few contaminant molecules in the purification could also be labelled with Alexa-568 dye during purification or some γ-TuRCs could exist as incomplete rings as shown recently (Liu et al., 2020). We have now reported this in the text in subsection “Contribution of end architecture of γ-TuRC to microtubule nucleation”, “At the highest tubulin concentrations, approximately 10-15% of γ-TuRCs nucleate MTs in the TIRF assays”, and how this calculation was made is detailed in the Materials and methods section.

[Editors’ note: what follows is the authors’ response to the second round of review.]

Summary:

The assembly and organization of the microtubule cytoskeleton requires nucleation of microtubules by the nucleator γ-TuRC, but the underlying mechanism has remained obscure. Recent Cryo-EM structures of γ-TuRC have raised additional questions by showing that native γ-TuRC is in a conformation that is not well suited to template microtubule nucleation. This study provides first insight by establishing TIRF imaging from purified, immobilized γ-TuRCs in vitro. Using a combination of biochemistry and modeling the study explains how an "imperfect" nucleation template can function as nucleation platform and identifies several crucial determinants of this activity. Rather than activation of γ-TuRC by a conformational switch prior to nucleation, the authors propose that the conformational switch occurs as a result of nucleation.

The authors have provided very thorough revisions, presenting and including a substantial amount of new data. The manuscript has been significantly improved and despite the lack of structural support data by EM in the manuscript, the main conclusions are well supported and we consider it now suitable for publication in eLife. There are, however, a few remaining issues that we would like the authors to address.

Essential revisions:

1) We appreciate that you have modified the title, but it now seems to not make complete sense. We would suggest "The transition state and regulation of γ-TuRC-mediated microtubule nucleation revealed by single molecule microscopy".

That sounds good and we have now edited the manuscript’s title as requested.

2) "Notably, the distinct minimal tubulin concentration needed for seeds to assemble a MT as compared to a previous report 24 results from the differences in assay conditions." The authors should be clearer here, explaining in more detail (as they did in the response to reviewers) that they got the same result as previous authors when they used their particular conditions.

We agree with this point and clarified this point further in the manuscript. In subsection “Contribution of end architecture of γ-TuRC to microtubule nucleation”, we state, “Notably, when this experiment was replicated with the coverslip preparation and assay conditions reported previously25, a high concentration of tubulin was necessary for seeds to assemble MTs in agreement with the previous work25. However, our assay conditions, that were used to compare seed-templated MT assembly with γ-TuRC-mediated nucleation side-by-side, result in a low, minimal tubulin concentration that is needed for seed-mediated MT assembly.”

3) "In summary, because γ-TuRC positions an array of γ-tubulins at its nucleation interface that are thought to stabilize intrinsically weak, lateral αβ/αβ-tubulin interaction 9,10,13,14,17,20,22147, MT nucleation by γ-TuRC has been proposed to function similar to polymerization of a MT end. Here we show several lines of evidence that γ-TuRC-mediated nucleation has distinct characteristics from MT polymerization and assembly from blunt MT seeds. While growth speed of MTs nucleated from γ-TuRC or templated from MT seeds is similar (Figure 3—figure supplement 1D), γ-TuRC molecules do not nucleate MTs at low tubulin concentration where MT polymerization can occur." It is important to point out here that this relates specifically to the purified γTuRCs that nucleate microtubules in the absence of other cellular factors. While the authors do show that some of the proposed activators do not activate the γ-TuRC in their assays, they cannot yet rule out that other proteins do function in vivo to promote the correct configuration of the γTuRC. They mention this in the Discussion section, but they are making the statement in the Results section, and it is a bit misleading to group all γTuRCs into this poor-nucleation bracket.

We appreciate this point by the reviewers. In subsection “Contribution of end architecture of γ-TuRC to microtubule nucleation”, we appended the specified text with, “While these results were obtained with endogenous γ-TuRCs purified from cytosol, it remains possible that specific factors at MTOCs can modulate γ-TuRC’s conformation and kinetics.”

4) Figure 6B, D – the authors state that there is no significant difference in nucleation rate when they add γTuNA or TPX2, but the graphs do show differences and no statistical test is provided.

We thank the reviewers for raising this important point. We used the 95% confidence interval to indicate the high degree of similarity. At the same time, we agree that our data can be represented in another way that quantitatively reports the small effect of TPX2 and γ-TuNA. In accordance with how the effect of activation factors was reported in previous and recent works (Consolati et al., 2020; Liu et al., 2020; Choi et al., 2010; Kollman et al., 2010 and others), we have now presented this data as fold change. Specifically, CDK5RAP2’s γ-TuNA motif γ-TuRC-mediated nucleation by 1.4(± 0.02) -fold at t = 180 seconds (mean ± std for n = 2 biological replicates). TPX2 increases the nucleation activity of γTuRC by 1.2 (± 0.3) -fold (mean ± std for n = 3 biological replicates) at t = 180 seconds. We have also reported the effect of XMAP215 in a similar fashion in text in subsection “XMAP215 promotes microtubule nucleation by strengthening the longitudinal bond energy between γ-TuRC and αβ-tubulin” for the corresponding Figure 7B. Further, we have edited the sub-headings of Figure 6B and Figure 6D to reflect these modifications.

5) "Recently, XMAP215 was discovered to be a nucleation factor that synergizes with γ-TuRC". There is a new pre-print from Trisha Davis' group showing that nucleation by γTuRC and XMAP215 is additive, not synergistic http://biorxiv.org/content/early/2020/05/23/2020.05.21.109561. It would be useful for the reader if the authors would discuss how this compares to their work here.

We appreciate this comment by the reviewers. We will summarize the recent work from Davis lab below and then explain how we will incorporate it.

As detailed below, we have several concerns regarding the pre-print on bioRxiv including that it is incomplete. First, in King et al., 2020, the data for supplementary figures has not been provided that is necessary to understand how “additive” and “synergistic” effects of XMAP215 with γ-tubulin filaments were calculated with a theoretical model. Little detail for this is provided in the Materials and methods section. As a result, it is unclear to us how the additive effect of XMAP215 was established, an understanding which is necessary to comment on this finding in our manuscript. Second, and most importantly, we note that the new preprint used turbidity measurement to assay MT assembly, which is not a direct measurement of MT nucleation. Turbidity combines the measurement of number of MTs (nucleation) with the total length of MTs. The latter is an important effect that cannot be ignored with polymerases like XMAP215, which increase MT length too. Instead, a direct nucleation assay is needed to clearly measure XMAP215’s effect, and distinguish between synergistic or additive scenarios. Finally, from the limited information provided in King et al., 2020, we gather that the turbidity curves reported are a normalized (not absolute) measurement of the total MT mass assembled in the individual reaction. The 10% assembly time from normalized curves was compared or likely summed across conditions to assign a role to XMAP215. We believe that instead of this analysis, quantitatively comparing the total MT mass (prior to normalization) will be necessary to assess if XMAP215 has an additive or synergistic effect.

Indeed, work from two other labs alongside our previous work have established a synergy between XMAP215 homologs with γ-tubulin complexes using varied model systems (Gunzelmann et al., 2018; Consolati et al., 2020; Thawani et al., 2018). In all three works, when measured with an assay where number of MTs were directly counted, the two molecules alone are inefficient in MT nucleation but have a synergistic effect. Finally, our current work alongside Thomas Surrey’s recent work (Consolati et al., 2020) further supports a synergistic effect with single molecule assays.

Nevertheless, we cite and give credit the work on bioRxiv and incorporate the reviewers’ suggestion as best as possible. In Subsection “Role of putative activation factors in γ-TuRC mediated nucleation”, we changed the introductory sentence to the following one, “ Recently, XMAP215 was discovered to be a nucleation factor that synergizes with γ-TuRC in X. laevis and S. cerevisiae29,50, or works in an additive manner with γ-tubulin51.” Then, we resolve this paradox at the end of the XMAP215 section on page16-17 with “Altogether, our results confirm that XMAP215 indeed functions synergistically with γ-TuRC, in agreement with recent works15,29,50. Most importantly, our results show that, while the transition state is defined by γ-TuRC’s conformation, XMAP215 strengthens the longitudinal γ-/αβ-tubulin bond to function as a bona-fide nucleation factor.”

6) The authors should discuss in more detail recently published work by the Surrey group (2020), which also shows, as the current manuscript, TIRF imaging of nucleation from immobilized γ-TuRCs. There are some differences e.g. regarding the critical number of dimers that need to assemble on gTuRC and regarding overall gTuRC activity in the assay. It would be very useful for the reader to know how the observations/conclusions in the two studies agree or differ.

We have expanded on our Discussion of this recently published work with the sentences:

“Notably, a parallel work also reported MT nucleation from single, human γ-TuRC molecules recently15. While the majority of findings agree with our work, 6.7 dimers were required in the critical nucleus and an overall lower activity of γ-TuRC (0.5%) was found15. Low structural integrity of purified γ-TuRC from incorporation of BFP-tagged GCP2 and a higher ratio of γ-tubulin sub-complexes, or species-specific variation in γ-TuRC properties could explain these differences.”

In the revised manuscript file, we have also made changes to the bibliography to reflect the final, published versions of the cryo-EM works.

https://doi.org/10.7554/eLife.54253.sa2

Article and author information

Author details

  1. Akanksha Thawani

    Department of Chemical and Biological Engineering, Princeton University, Princeton, United States
    Contribution
    Conceptualization, Software, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4168-128X
  2. Michael J Rale

    Department of Molecular Biology, Princeton University, Princeton, United States
    Contribution
    Investigation, Purified NME7 and CDK5RAP2 fragments and contributed to the related experiments
    Competing interests
    No competing interests declared
  3. Nicolas Coudray

    Department of Cell Biology, New York University School of Medicine, New York, United States
    Contribution
    Methodology, Provided advice on EM reconstruction of αβ-tubulin filaments
    Competing interests
    No competing interests declared
  4. Gira Bhabha

    Department of Cell Biology, New York University School of Medicine, New York, United States
    Contribution
    Methodology, Provided advice on EM reconstruction of αβ-tubulin filaments
    Competing interests
    No competing interests declared
  5. Howard A Stone

    Department of Mechanical and Aerospace Engineering, Princeton University, Princeton, United States
    Contribution
    Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9670-0639
  6. Joshua W Shaevitz

    1. Lewis-Sigler Institute for Integrative Genomics, Princeton, United States
    2. Department of Physics, Princeton University, Princeton, United States
    Contribution
    Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8809-4723
  7. Sabine Petry

    Department of Molecular Biology, Princeton University, Princeton, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Project administration, Writing - review and editing
    For correspondence
    spetry@Princeton.EDU
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8537-9763

Funding

American Heart Association (17PRE33660328)

  • Akanksha Thawani

Princeton University (Charlotte Elizabeth Procter Honorific Fellowship)

  • Akanksha Thawani

Howard Hughes Medical Institute (Gilliam fellowship)

  • Michael J Rale

National Science Foundation (Graduate Student Fellowship)

  • Michael J Rale

National Institute of General Medical Sciences (R00GM112982)

  • Gira Bhabha

National Institute of General Medical Sciences (1DP2GM123493)

  • Sabine Petry

Pew Charitable Trusts (00027340)

  • Sabine Petry

David and Lucile Packard Foundation (2014-40376)

  • Sabine Petry

National Science Foundation (PHY-1734030)

  • Joshua W Shaevitz

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Brian Mahon, Brianna Romer and Sophie Travis for advice on collecting and processing of electron microscopy data, as well as Petry lab members for discussions. We thank David Agard and Michelle Moritz for sharing unpublished data and for discussions. This work was supported by an American Heart Association predoctoral fellowship 17PRE33660328 and a Princeton University Honorific Fellowship (both to AT), a Howard Hughes Medical Institute Gilliam fellowship and a National Science Foundation graduate research fellowship (both to MJR), NIGMS R00GM112982 (to GB), NIH New Innovator Award 1DP2GM123493, Pew Scholars Program in the Biomedical Sciences 00027340, David and Lucile Packard Foundation 2014–40376 (all to SP), and the Center for the Physics of Biological Function sponsored by the National Science Foundation grant PHY-1734030.

Ethics

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to approved Institutional Animal Care and Use Committee (IACUC) protocol # 1941-16 of Princeton University.

Senior Editor

  1. Anna Akhmanova, Utrecht University, Netherlands

Reviewing Editor

  1. Jens Lüders, Institute for Research in Biomedicine, Spain

Reviewer

  1. Jens Lüders, Institute for Research in Biomedicine, Spain

Publication history

  1. Received: December 7, 2019
  2. Accepted: June 15, 2020
  3. Accepted Manuscript published: June 15, 2020 (version 1)
  4. Version of Record published: July 6, 2020 (version 2)
  5. Version of Record updated: July 9, 2020 (version 3)

Copyright

© 2020, Thawani et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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