1. Structural Biology and Molecular Biophysics
  2. Cell Biology
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A mutation uncouples the tubulin conformational and GTPase cycles, revealing allosteric control of microtubule dynamics

  1. Elisabeth A Geyer
  2. Alexander Burns
  3. Beth A Lalonde
  4. Xuecheng Ye
  5. Felipe-Andres Piedra
  6. Tim C Huffaker
  7. Luke M Rice  Is a corresponding author
  1. University of Texas Southwestern Medical Center, United States
  2. Cornell University, United States
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Cite this article as: eLife 2015;4:e10113 doi: 10.7554/eLife.10113

Abstract

Microtubule dynamic instability depends on the GTPase activity of the polymerizing αβ-tubulin subunits, which cycle through at least three distinct conformations as they move into and out of microtubules. How this conformational cycle contributes to microtubule growing, shrinking, and switching remains unknown. Here, we report that a buried mutation in αβ-tubulin yields microtubules with dramatically reduced shrinking rate and catastrophe frequency. The mutation causes these effects by suppressing a conformational change that normally occurs in response to GTP hydrolysis in the lattice, without detectably changing the conformation of unpolymerized αβ-tubulin. Thus, the mutation weakens the coupling between the conformational and GTPase cycles of αβ-tubulin. By showing that the mutation predominantly affects post-GTPase conformational and dynamic properties of microtubules, our data reveal that the strength of the allosteric response to GDP in the lattice dictates the frequency of catastrophe and the severity of rapid shrinking.

https://doi.org/10.7554/eLife.10113.001

eLife digest

Protein filaments called microtubules help move cargo around inside cells. Chromosomes, which contain the cell’s genetic blueprints, are the microtubule’s most precious cargo. Before a cell divides, microtubules grow from the ends of the dividing cell towards the middle, where they attach to the chromosomes that are lined up along the centerline. Then the microtubules shrink and drag the chromosomes back to the opposite ends of the cell. This allows each of the new cells to get one copy of each chromosome.

When the microtubules are growing, a molecule called guanosine triphosphate (or GTP) is attached to the proteins at the end of the filament. This acts like a cap and protects the microtubule from shrinking. Later a chemical reaction converts GTP into GDP (short for guanosine diphosphate). Without the protective GTP cap, the microtubule quickly shrinks. At the same time, the proteins that make up the microtubule also change shape. In the microtubule, the proteins adopt a straight shape when GTP is attached. The proteins favor a different shape in the microtubule when GDP is attached. However, it is unclear if or how these shape changes contribute to how a microtubule grows or shrinks.

Geyer et al. now show how this shape shifting can influence microtubule shrinking, by first identifying a mutation in yeast microtubule proteins that cause the proteins to remain straight even when GDP is attached. Next, powerful microscopes were used to make time-lapse videos of the mutated microtubules. This allowed Geyer et al. to observe how the mutated microtubules behaved and compare this to the behavior of normal microtubules.

The experiments revealed that the mutated microtubules were less likely to begin shrinking than typical microtubules. The mutated microtubules also shrunk more slowly. These findings indicate that the shape changes control the speed of shrinking and frequency of entering the shrinking phase. These new details about the control of microtubule growth and shrinkage may help scientists studying how cell division happens in both healthy and cancerous cells.

https://doi.org/10.7554/eLife.10113.002

Introduction

Microtubules are dynamic polymers of αβ-tubulin that undergo GTPase-dependent switching between phases of growing and rapid shrinking (reviewed in Desai and Mitchison, 1997). Microtubule growing and shrinking rates, and switching frequencies, are necessary for proper function, highly regulated by cellular factors (Howard and Hyman, 2003), and targeted by widely used anti-mitotic anticancer drugs (Jordan and Wilson, 2004; Amos, 2011). αβ-tubulin also undergoes a conformational cycle during polymerization and depolymerization (Mandelkow et al., 1991a; Chrétien et al., 1995) Unpolymerized αβ-tubulin adopts a 'curved' conformation (Ravelli et al., 2004; Rice et al., 2008; Buey et al., 2006) that is not compatible with the 'straight' microtubule lattice (Nogales et al., 1998). In the body of the microtubule, αβ-tubulin can adopt distinct ‘expanded’ or ‘compacted’ straight conformations depending on nucleotide state (Alushin et al., 2014; Hyman et al., 1995). Partially straightened conformations of αβ-tubulin are thought to occur as polymerization intermediates on the microtubule end.

Recent studies have begun to reveal how microtubule regulatory proteins take advantage of these different conformations. In one well-characterized example, end-binding proteins in the EB1 family have been shown to mark the plus-end cap of microtubules by binding preferentially to the ‘expanded’, GTP-like conformation of αβ-tubulin that only occurs near the end of the growing microtubule lattice (Maurer et al., 2011; 2012; Zanic et al., 2009; Alushin et al., 2014). Another recent example of selective interactions with specific conformations of αβ-tubulin comes from work in our laboratory showing that TOG domains from the microtubule polymerase Stu2p bind preferentially to the curved conformation of αβ-tubulin (Ayaz et al., 2012; 2014), and that this preference of TOG domains for curved αβ-tubulin is probably what allows the polymerase to localize to the growing tip of the microtubule (Ayaz et al., 2014). These and numerous studies of other regulatory proteins (Desai et al., 1999; Gigant et al., 2000; Peters et al., 2010; Alushin et al., 2010; Bechstedt et al., 2014) contribute to the emerging view that selective binding to distinct conformations of αβ-tubulin represents a common and important strategy for recognizing and controlling microtubules.

How the αβ-tubulin conformational cycle contributes to microtubule dynamics is less understood. Indeed, even though structural rearrangements in αβ-tubulin have long been known to occur during and after microtubule incorporation (Mandelkow et al., 1991b; Chrétien et al., 1995; Hyman et al., 1995), actually showing that these rearrangements contribute to and are required for the dynamic properties of microtubules remains challenging (Rice et al., 2008; Wang and Nogales, 2005; Buey et al., 2006; Jánosi et al., 2002; Molodtsov et al., 2005). For example, GTP-bound αβ-tubulin ‘straightens’ during the formation of longitudinal and lateral lattice contacts (Buey et al., 2006; Rice et al., 2008; Nawrotek et al., 2011), but how (or even if) the energetic cost of these assembly-dependent conformational changes affects polymerization dynamics remains debated. A similar argument can be made about the αβ-tubulin conformational changes that occur as a consequence of GTP hydrolysis deeper in the microtubule lattice. One reason these questions are still debated is that to date in vitro microtubule dynamics have only been measured for a small number of αβ-tubulin mutants (Gupta et al., 2002; Dougherty et al., 2001; Sage et al., 1995), and it has not yet been possible to link altered dynamics to a change in the αβ-tubulin conformational cycle.

In the present study we sought to discover how multiple αβ-tubulin conformations affect microtubule dynamics by identifying mutations that selectively perturb some aspect of the conformational cycle. We reasoned that mutating buried residues with different packing environments in the curved and straight conformations might provide a way to perturb the conformational cycle. We focused on helix H7 because it is an element that is positioned differently in the ‘curved’ (unpolymerized) and ‘straight’ (polymerized) conformations of αβ-tubulin (Nogales et al., 1998; Ravelli et al., 2004)(Figure 1A), and it may act as a structural relay connecting the ‘top’ and ‘bottom’ of the tubulin subunits (Amos, 2004). The equivalent helix in α-tubulin was also recently observed to move in response to GTP hydrolysis in the lattice (Alushin et al., 2014). We chose to study the T238A mutation in β-tubulin because this sidechain resides on helix H7 and is buried in the core of the protein (Figure 1A), and because prior studies (Thomas et al., 1985; Machin et al., 1995) showed that the β:T238A phenotype was consistent with hyperstable microtubules. The molecular mechanism by which the β:T238A mutation affects microtubule dynamics is unknown, in part because the polymerization dynamics of β:T238A αβ-tubulin have not been studied in vivo or in vitro. Our studies now reveal that the β:T238A mutation affects microtubule dynamics by dramatically reducing the frequency of catastrophe and the rate of shrinking, and that the mutation affects microtubule structure by attenuating post-GTPase conformational changes in the lattice. The effects of the mutation are reminiscent of the way the microtubule stabilizing drug taxol affects microtubule structure (Alushin et al., 2014). By showing that the β:T238A mutation weakens the coupling between the conformational and nucleotide cycles, our results demonstrate that catastrophe can occur despite diminished post-GTPase conformational changes in the microtubule lattice, and that the strength of the allosteric response to GDP dictates the rate of microtubule shrinking and the frequency of microtubule catastrophe.

αβ-tubulin containing a buried mutation in β-tubulin gives hyperstable microtubules in vivo and in vitro.

(A) Superposition of polymerized (‘straight’, green, PDB 1JFF) and unpolymerized (‘curved’, blue, PDB 4FFB) conformations of β-tubulin. T238 is solvent inaccessible and resides on a helix (H7) that undergoes a piston-like movement between the two conformations. The view of β-tubulin shown here is as if from the center of the microtubule looking out, with the plus end at the top. (B) Time-lapse imaging of live yeast shows that cells expressing T238A αβ-tubulin make longer, less dynamic microtubules than their wild-type (WT) counterparts. N = 13, 18 for WT and T238A microtubules respectively, with t = 35, 72 min total time observed. See also Videos 1,2. Bar = 2 µm (C) Negative stain electron micrographs (magnification: 23,000x) of wild-type (left) and β:T238A (right) microtubules. Mutant microtubules have normal structure. Bar = 100 nm. (D) In vitro, T238A and wild-type microtubules show similar concentration-dependent growth rates (slopes: 29.6 +/- 1.8 µm/hr/µM for wild-type, 25.5 +/- 0.8 µm/hr/µM for β:T238A, the differences between the slopes are not statistically significant; x-intercepts: 0.12 µM for wild-type, 0.033 µM for β:T238A, this difference is statistically significant; significance of differences in regression parameters was evaluated using GraphPad Prism). Representative kymographs are shown above the plots. (N = 12 for all points except 0.2 µM where N = 20; bars show s.e.m.) (E) T238A microtubules catastrophe less frequently than wild-type (top; N = 99, 115 for wild-type and T238A respectively, bars show s.d.) and show a ∼hundredfold slower rate of post-catastrophe shrinking (bottom; N = 16 for wild-type and T238A, bars show s.e.m.).

https://doi.org/10.7554/eLife.10113.003

Results

The buried T238A mutation in β-tubulin hyperstablizes microtubules in vivo and in vitro

The β:T238A mutation was previously identified by virtue of drug- and temperature-sensitive phenotypes that were consistent with hyperstable microtubules (Thomas et al., 1985). However, prior studies of this mutant have not directly examined its polymerization dynamics in vivo or in vitro (Machin et al., 1995; Dorn et al., 2005). To obtain insight into how the mutation affects polymerization dynamics in cells, we used time lapse imaging in GFP-Tub1p expressing strains to measure microtubule dynamics of wild-type and β:T238A-containing yeast. These measurements revealed that β:T238A-tubulin forms static microtubules (neither growing nor shrinking) that are on average over 50% longer than the dynamic microtubules that form in a strain with wild-type β-tubulin (Figure 1B and Videos 1,2). Because the mutated site is solvent inaccessible, this striking change in microtubule dynamics cannot be the result of a direct perturbation of a polymerization interface or of an interaction with one or more regulatory proteins.

Video 1
Microtubule dynamics in wild type yeast.

Time-lapse images of Tub1-GFP in TUB2 (β-tubulin) cells were taken at 15-s intervals; video plays at 4 frames/s. A frame from this video is shown in Figure 1B.

https://doi.org/10.7554/eLife.10113.004
Video 2
Microtubule dynamics in tub2-150 (T238A mutation in Tub2p) yeast.

Time-lapse images of Tub1-GFP in tub2-150 cells were taken at 15-s intervals; video plays at 4 frames/s. A frame from this video is shown in Figure 1B.

https://doi.org/10.7554/eLife.10113.005

To determine how the buried β:T238A mutation affected microtubule dynamics in vitro, we purified β:T238A αβ-tubulin from an overexpressing strain of yeast (Johnson et al., 2011) and used time-lapse differential interference contrast microscopy to measure its polymerization dynamics. We were unable to measure mutant and wild-type microtubule dynamics at equivalent concentrations, because β:T238A αβ-tubulin showed abundant spontaneous nucleation at the higher concentrations where we measured wild-type, and wild-type αβ-tubulin does not elongate measurably at the low concentrations where we were able to measure β:T238A dynamics without excessive nucleation. Mutant and wild-type microtubules nevertheless show similar concentration-dependent elongation rates: fitting lines to mutant and wild-type data reveals that the x-intercepts of the two datasets (0.12 and 0.033 µM for wild-type and β:T238A, respectively) differ by a factor of ∼3.5 and that the difference in slope (29.6 and 25.5 µm/hr/µM for wild-type and β:T238A, respectively) is not statistically significant (Figure 1D). Because the x-intercept and slope respectively relate to the apparent affinity and association rate constant for elongation, our data indicate that the mutation has little effect on the apparent biochemistry of microtubule elongation. Consistent with this biochemical similarity, negative stain electron microscopy revealed that mutant and wild-type microtubules show similar structure (Figure 1C). In striking contrast to the shared elongation behavior, after catastrophe β:T238A microtubules shrink roughly hundredfold more slowly than wild-type (1.1 µm/min for β:T238A compared to 96 µm/min for wild-type, Figure 1E, bottom). Thus, the mutation significantly strengthens the lattice contacts that dictate the rate of microtubule shrinking. Finally, β:T238A microtubules also undergo catastrophe much less frequently than wild-type. The lower catastrophe frequency we observed is especially notable when considering that in these assays the T238A microtubules were growing much slower than wild-type because of the ∼threefold lower concentration of αβ-tubulin used for the mutant (Figure 1E, top).

Mutant-induced changes in polymerization dynamics do not result from defective GTPase activity

The β:T238A mutation stimulated spontaneous nucleation and reduced the frequency of catastrophe and the rate of shrinking, all without substantially affecting elongation. It seemed possible that a defective GTPase cycle might explain these observations. We reasoned that if the increased spontaneous nucleation of the β:T238A mutant resulted from slower/defective GTPase activity, then both mutant and wild-type should nucleate with similar efficiency when GTP hydrolysis cannot occur. We initially attempted to use GMPCPP, the hydrolysis-resistant nucleotide of choice for vertebrate microtubules (Hyman et al., 1992), but GMPCPP did not support elongation of yeast microtubules in our dynamics assays. Yeast microtubules polymerized readily in the presence of GTPγS, however, indicating that GTPγS better mimics GTP for yeast microtubules. We observed that even in the presence of GTPγS, wild-type microtubules show substantially less nucleation than T238A microtubules (Figure 2A,B). Thus, the abundant nucleation from the mutant cannot be ascribed to a defect in GTPase activity. Instead, the mutation must be affecting some other property that limits spontaneous nucleation in wild-type αβ-tubulin.

T238A αβ-tubulin undergoes spontaneous nucleation more readily than WT, even in the presence of a nonhydrolyzable GTP analog, GTPγS.

(A) Fluorescent images of crosslinked microtubules from spontaneous nucleation reactions. Even at low concentrations and in the presence of GTPγS, T238A tubulin shows increased spontaneous nucleation compared to WT. GTPγS reactions are presented next to each other to facilitate a side-by-side comparison. Scale bar in top left is 5 µm. (B) Microtubule spindown reactions show that under the same concentration range, in the presence of GTPγS, T238A tubulin produces a greater proportion of microtubules which sediment into the pellet. Gel images of supernatant and pellet fractions (top). (C) T238A microtubules do not accumulate GTP or GDP.Pi compared to wild-type. Images show TLC analysis of exchangeable nucleotide content of microtubules grown with GTP or GTPγS. Microtubules were spontaneously assembled using higher concentrations than for the dynamics assays: wild-type microtubules were prepared at 2 µM with GTP and 1 µM with GTPγS. β:T238A microtubules were prepared at 1 µM with either nucleotide. (D) Quantification of TLC data, n = 3 and errors shown represent s.d. *indicates one condition where we could not detect any Pi in one of the replicates; instead of using 0 we used the lower of the two other trials.

https://doi.org/10.7554/eLife.10113.006

To support the idea that the mutation-induced changes in dynamics result from something other than a defect in GTPase activity, we assayed the nucleotide content of wild-type and β:T238A microtubules. We allowed wild-type and β:T238A αβ-tubulin to spontaneously polymerize at 1 or 2 µM concentration (within the range of concentrations tested in Figure 2A) in the presence of 32P-GTP. We harvested the microtubules by centrifugation, denatured them to release bound nucleotides, and analyzed the nucleotide content at the exchangeable site using thin layer chromatography (TLC) (Figure 2C,D). We probed for a GTPase defect using α-labeled nucleotide to measure the amounts of GTP and GDP in microtubules, and for a phosphate release defect using γ-labeled nucleotide to measure the amounts of GTP and Pi in microtubules. To avoid possible complications that might arise from the very different catastrophe frequencies and shrinking rates, we initially performed these assays under ‘catastrophe free’ and slow shrinking conditions by using (unlabeled) GTPγS to support assembly. These experiments revealed that wild-type and mutant microtubules contain similarly low amounts of GTP (2–4% of total exchangeable nucleotide) and Pi (fewer than 3% of exchangeable sites) (Figure 2C,D). When we performed these assays using (unlabeled) GTP to support microtubule assembly, β:T238A microtubules contained similarly low amounts of GTP and Pi as we observed with GTPγS but wild-type microtubules appeared to contain a substantially greater amount (∼50%) of GTP (Figure 2C,D). Yeast microtubules have previously been reported to contain more GTP than vertebrate microtubules (Dougherty et al., 1998). In control experiments (not shown) we confirmed that vertebrate microtubules contain very little GTP and that individual heterodimers were not pelleting. Because we observed very fast shrinking (Figure 1), little 32P-GTP in microtubules grown with GTPγS (Figure 2C), and normal plus-end recognition by an EB1 family protein (see below), it seems unlikely that wild-type yeast microtubules contain substantial amounts of GTP. The GTP containing material in our assay might instead reflect oligomers that form more readily for yeast αβ-tubulin. Whatever the mechanism, these experiments demonstrate that β:T238A αβ-tubulin remains competent to hydrolyze GTP, and that the altered nucleation and shrinking behavior cannot be ascribed to an accumulation of GTP or GDP.Pi that might result from a defect in assembly-dependent GTPase activity or phosphate release. Some other, nonenzymatic mechanism must therefore account for the observed changes in polymerization dynamics.

The buried T238A mutation in β-tubulin does not detectably ‘straighten’ unoligomerized αβ-tubulin

Unoligomerized GTP-bound αβ-tubulin adopts a curved conformation that is not compatible with the straight microtubule lattice (Nawrotek et al., 2011; Pecqueur et al., 2012; Ayaz et al., 2012; 2014). It seemed possible that the β:T238A mutation might stabilize microtubules by shifting the conformational preference of unpolymerized αβ-tubulin to favor a straight(er) conformation that is more compatible with the microtubule lattice. We used quantitative TOG binding assays (Ayaz et al., 2014) to determine if the β:T238A mutation changed the ‘curvature’ of unpolymerized αβ-tubulin. The rationale for this approach is based on the fact that TOG domains bind tightly to curved αβ-tubulin but very weakly to straight (Figure 3A) (Ayaz et al., 2012; 2014 ). Accordingly, β:T238A αβ-tubulin should bind less tightly to a TOG domain if the β:T238A mutation appreciably changes the conformation of αβ-tubulin, for example by straightening it. TOG1 and TOG2, two different TOG domains from the yeast microtubule regulatory factor Stu2p (Wang and Huffaker, 1997), show very similar affinity (within a factor of two) for unpolymerized T238A αβ-tubulin as they do for wild-type (Figure 3B). In energetic terms, this modest difference in affinity is roughly equivalent to one hydrogen bond. While we cannot state that the mutant and wild-type αβ-tubulin adopt identical conformations, their very similar TOG binding properties indicate that any differences in conformation must be small. Thus, we conclude that the β:T238A mutation does not significantly change the conformation of unpolymerized αβ-tubulin.

The buried β:T238A does not appreciably straighten unpolymerized αβ-tubulin.

(A) Cartoon schematics illustrating that TOG domains (blue) bind tightly to curved αβ-tubulin (pink and green; left) but weakly to straight αβ-tubulin (right). (B) Isotherms for individual Stu2 TOG domains binding to T238A αβ-tubulin. Both TOG domains bind with comparable affinity to T238A and WT tubulin. 1 Σσ confidence intervals for fitted affinities are provided in parentheses.

https://doi.org/10.7554/eLife.10113.007

The β:T238A mutation suppresses GTPase-dependent conformational changes in the microtubule lattice

Recent cryo-EM studies of microtubule structure provided atomic models for distinct microtubule lattices containing GTP, GTPγS, or GDP, with GTP favoring an ‘expanded’ form and GTPγS and GDP favoring a ‘compacted’ form (Alushin et al., 2014; Zhang et al., 2015). Plus-end tracking proteins in the EB1 family have been shown to discriminate between these GTP-like and GDP-like lattices (Maurer et al., 2011; 2012; 2014; Zanic et al., 2009). We used Bim1p (Schwartz et al., 1997), the yeast EB1 protein, to investigate if the β:T238A mutation affected αβ-tubulin conformation in the lattice.

Control experiments revealed that a Bim1-GFP fusion protein tracked the growing end of yeast microtubules, and that Bim1-GFP coated the entire length of GTPγS-containing yeast microtubules with cap-like intensity (Figure 4). We therefore infer that like other EB1 proteins on vertebrate microtubules (Zanic et al., 2009; Maurer et al., 2011), Bim1 discriminates between GTP and GDP forms of the yeast microtubule lattice (presumably, expanded and compacted). In marked contrast to its behavior on wild-type microtubules, at 50 nM concentration Bim1-GFP coated the entire length of ‘dynamic’ β:T238A microtubules with cap-like intensity. At lower concentrations (5 nM), the Bim1-GFP coat on the body of β:T238A microtubules is slightly weaker than at the cap but still substantially more intense than on wild-type (Figure 4B). This stronger coating reflects tighter Bim1 binding to mutant microtubules, and it occurs in spite of the fact that the mutant microtubules contain very little GTP or GDP.Pi in the exchangeable site (Figure 2). Thus, the β:T238A mutation attenuates the conformational response to GTP hydrolysis in the microtubule lattice. The structural consequence of the mutation resembles the effect of taxol binding, which also promotes an expanded conformation of αβ-tubulin in a GDP lattice (Alushin et al., 2014). Simply put, the mutation appears to have substantially uncoupled the conformational cycle from the nucleotide cycle in the lattice, allowing αβ-tubulin to retain GTP-lattice-like character even in a GDP lattice.

The buried β:T238A attenuates the conformational response to GDP in the lattice.

(A) Cartoon schematics illustrating the basis of the assay: EB1 proteins (blue outlined ovals) bind tightly to the GTP-lattice conformation of αβ-tubulin (left) but weakly to the GDP-like conformation (right). (B) Images of the distribution of Bim1-GFP (an EB1 family protein) on wild-type (left and center) or mutant microtubules (right). 4 microtubules are shown for each condition, and Bim1-GFP was present at 50 nM and 5 nM concentration for all microtubules. Plots represent average Bim1-GFP intensity on n = 9 microtubules for each condition. Error bars represent s.e.m. Cartoons illustrate the likely lattice conformation inferred from the Bim1-GFP binding. β:T238A microtubules appear GTP-like even though they contain GDP. Scale bars: 1 μm.

https://doi.org/10.7554/eLife.10113.008

The β:T238A mutation affects αβ-tubulin curvature on the microtubule end

The Bim1 experiments described above do not report on the conformation of αβ-tubulin at the very microtubule end because EB1 proteins do not bind there (Maurer et al., 2012; 2014). After fortuitously discovering that isolated TOG domains from the microtubule polymerase Stu2p stimulate the depolymerization of stabilized microtubules in a dose-dependent way, it seemed that TOG domains might provide an alternative way to probe the conformation of αβ-tubulin on the microtubule end. In light of the fact these TOG domains bind preferentially to the curved conformation of αβ-tubulin, we reasoned that the underlying cause of this induced depolymerization was TOG-mediated stabilization of the curved and faster dissociating conformation of αβ-tubulin on the microtubule end (Figure 5). According to this view, TOG-induced depolymerization thus provides an assay that probes the linkage (or lack thereof) between changes in αβ-tubulin conformation on the polymer end and microtubule shrinking rate.

Using drug-stabilized microtubules as a substrate, the rate of TOG-induced depolymerization increases linearly over at least a 20-fold range of TOG concentration. This linear concentration-dependence is consistent with a collisional mechanism in which the rate-limiting step is a TOG domain arriving at the microtubule end. It is not consistent with a mechanism in which the rate-limiting step is some kind of slow αβ-tubulin conformational change. Drug-stabilized β:T238A microtubules depolymerize substantially slower than wild-type at the same concentration of TOG domain (Figure 5). Only at higher TOG concentrations do β:T238A microtubules show appreciable dose-dependent TOG-induced depolymerization (Figure 5). Similar results were obtained using MTs stabilized with GTPγS (Figure 5).

β:T238A microtubules are resistant to TOG-induced depolymerization.

(A) A time series of images of stabilized fluorescent microtubules show that under the same concentration of TOG1, T238A microtubules depolymerize substantially slower than WT microtubules. Scale bars: 1 µm. (B) Quantification of the dose-dependence of the rate of TOG-induced depolymerization. Inset: a plot showing that under higher TOG1 concentrations, T238A microtubules also undergo induced depolymerization. We observed similar rates of TOG-induced depolymerization using DIC instead of fluorescence to monitor microtubule length (black triangles), as well as using GTPγS instead of epothilone as a stabilizing reagent (orange diamonds, purple squares). Additional control experiments demonstrate that the TOG-induced depolymerization is greatly reduced when a weakly-binding TOG1 mutant (R200A) is used (green triangles). N = 30 for fluorescence measurements of wild-type TOG1-induced depolymerization of wild-type or β:T238A microtubules, N = 20 for the rest. Error bars represent s.e.m. (C) Cartoon model illustrating the mechanism of TOG-induced depolymerization, resulting from TOG-stabilization of tubulin subunits sampling curved conformation at the end of the microtubule. T238A microtubules depolymerize slower due to a decrease in tubulin subunits sampling the curved conformation once incorporated into the polymer.

https://doi.org/10.7554/eLife.10113.009

TOG domains bind with comparable affinity to curved wild-type and mutant αβ-tubulin, so differences in TOG affinity cannot explain the markedly slower TOG-induced shrinking of β:T238A microtubules. Differences in the amount of curved αβ-tubulin at wild-type and mutant microtubule ends could explain the observed differences in TOG-induced shrinking. In principle there might be other mechanisms that strengthen association without affecting the propensity to be curved on the microtubule end. However, while such ‘curvature invariant’ mechanisms in αβ-tubulin might explain the slow post-catastrophe shrinking of the mutant, the conformation-selective nature of TOG:tubulin interactions means that this alternative view is not easily reconciled with the observed differences in TOG-induced depolymerization. Thus, a mutant-induced change in the propensity to become/remain straight on the microtubule end is the simplest way to explain both the slower post-catastrophe shrinking and weaker TOG-induced depolymerization for the mutant. That the conformation of end-bound αβ-tubulins is affected by the β:T238A mutation is also consistent with the Bim1 coating, which demonstrated that the mutation affected αβ-tubulin conformation elsewhere in the lattice.

What is the structural origin of the β:T238A effects?

To gain insight into the local interactions responsible for the β:T238A effects on microtubule dynamics, we measured benomyl sensitivity/resistance and in vivo microtubule dynamics for serine and valine substitutions at the same position (Figure 6). Yeast expressing β:T238S αβ-tubulin are comparably benomyl resistant to β:T238A, but significantly less benomyl dependent (Figure 6A). By contrast, yeast expressing β:T238V αβ-tubulin are substantially less resistant to benomyl, and show no benomyl dependence (Figure 6A). Measurements of microtubule dynamics in vivo are consistent with the benomyl phenotypes: β:T238S microtubules are β:T238A-like (static) whereas β:T238V microtubules are more wild-type-like (dynamic but with somewhat slower shrinking) (Figure 6B). The trend with sidechain size and the strong effect obtained from the Thr to Ser mutation at position 238 suggest that some form of ‘steric overpacking’, not hydrogen bonding, contributes to destabilize the straight, expanded conformation of αβ-tubulin in microtubules containing GDP.

Insights into the mechanism underlying the β:T238A effects on the conformational cycle.

(A) Yeast with different substitutions for T238 (for each mutant, strains with and without GFP-Tub1p are shown) show intermediate degrees of benomyl resistance/dependence. The volume of the packing defect may be related to the magnitude of the resulting phenotype: T238S has a phenotype closer to T238A whereas T238V has a phenotype closer to wild type. (B) In vivo microtubule dynamics of yeast containing T238S and T238V αβ-tubulin. N = 24, 14 for T238S and T238V microtubules respectively, with t = 120, 43 min total time observed. (C) The buried β:C354A mutation immediately proximal to T238 (left) has also been shown to give slowly shrinking microtubules, and also shows an expanded lattice (right). The view of the structure is as if from the center of the microtubule looking out, with the plus end at the top. Scale bar: 1 µm.

https://doi.org/10.7554/eLife.10113.010

Reducing the size of β:C354, a buried sidechain that packs against β:T238 (Figure 6C), has also been shown to stabilize microtubules (Gupta et al., 2001; 2002). Indeed, β:C354A or β:C354S substitutions dramatically reduced the rate of microtubule shrinking and the frequency of catastrophe (Gupta et al., 2002). If these volume-reducing mutations also stabilize the expanded conformation of αβ-tubulin in a GDP lattice, it would provide complementary support for our overpacking-based rationale for the β:T238(A, S, V) effects. We purified β:C354A αβ-tubulin and used Bim1-GFP to probe its conformation in microtubules. β:C354A microtubules showed Bim1-GFP coating similar to what we observed for β:T238A, indicating the two mutations stabilize microtubules through a common conformational mechanism (Figure 6C). Positions 238 and 354 in β-tubulin show strong evolutionarily conservation (yeast numbering; β:238 is 65% T and 34% C whereas β:354 is 99% C), consistent with an important functional role for these residues. Together, the β:T238A and β:C354A results suggest that changes in packing volume near helix H7 can dictate the dynamic properties of microtubules by tuning an allosteric response to GDP.

Discussion

We demonstrate that relatively conservative buried mutations in β-tubulin can diminish the conformational response to GDP in the lattice, and that this altered structural response yields dramatic effects on microtubule dynamics. By discovering that mutations can uncouple the conformational and nucleotide cycles of αβ-tubulin, our findings provide new insights into ways that the αβ-tubulin conformational cycle dictates microtubule polymerization dynamics (Figure 7).

The αβ-tubulin conformational cycle and its impact on microtubule dynamics.

(A) Unoligomerized wild-type and β:T238A αβ-tubulin both adopt the curved conformation. That the straight conformation is strained (flare marks) even with the mutation-induced reduction in packing volume (cartooned by larger and smaller red circles for wild-type and β:T238A, respectively) suggests that extrinsic factors like interactions with the lattice control straightening. (B) Compared to wild-type, β:T238A αβ-tubulin is better able to populate a straight conformation on the end of GDP containing microtubules. ‘D’ indicates GDP underneath the bolded terminal subunit, compaction is represented as in Figure 4, and the white arrow cartoons trans-acting nucleotide (see text). The increased ability of the mutant to be straight could result from an altered response to GDP on the longitudinal interface, from more favorable interactions with the expanded lattice, or from a combination of both. (C) The schematic phase diagram illustrates that the mutation-induced changes to the αβ-tubulin conformational cycle decrease the threshold concentrations for appreciable elongation against catastrophe (line 1) and for spontaneous nucleation (line 2), and also narrows the gap between them. The normal αβ-tubulin conformational cycle contributes to microtubule dynamics and makes them more amenable to regulation.

https://doi.org/10.7554/eLife.10113.011

Our experiments using the EB1 family protein Bim1 showed that in the lattice a major effect of the β:T238A mutation is to substantially suppress conformational changes that normally occur as a consequence of GTPase activity (Hyman et al., 1995; Alushin et al., 2014; Zhang et al., 2015). The different responses of yeast and vertebrate microtubules to GMPCPP, and the lack of high-resolution structures for wild-type or mutant yeast microtubules, make it difficult to know the specific conformations involved and how they relate to the nucleotide cycle. However, the stronger binding of Bim1 to GDP-containing β:T238A microtubules indicates that the mutant microtubule lattice retains more GTP-like character than does wild-type. This observation about αβ-tubulin in the body of the microtubule cannot by itself explain the slow shrinking, however, because the rate of shrinking is determined by the properties of incompletely surrounded terminal subunits, on which EB1 does not report. Indeed, understanding how different lattice structures relate to dynamic properties of microtubules remains a significant challenge. Our experiments with TOG domains indirectly probed the conformation of terminal subunits and showed that the β:T238A mutation reduces the propensity of terminal αβ-tubulins to be curved. Thus, the slow shrinking and lower frequency of catastrophe in the mutant most likely result from the attenuated allosteric response to GDP; this in turn results in a straighter and more strongly associated GTP-like conformation on the microtubule end, despite having GDP at the longitudinal interface (Figure 7B).

It is remarkable that the majority of the mutation-induced effects are confined to post-GTPase conformational changes in the lattice. Not observing significant effects on elongation is somewhat surprising, because it had been anticipated that the conformational changes required to enter the lattice would contribute to polymerization dynamics by opposing incorporation into the lattice. The lack of substantial change in the observed concentration-dependence of elongation rates could indicate that the β:T238A mutations do not affect the propensity for αβ-tubulin to ‘straighten’ on a GTP lattice. However, this is not consistent with increased spontaneous nucleation of β:T238A αβ-tubulin in the presence of GTPγS, which indicated that the mutation did affect conformational transitions that occur in/on a GTP lattice. It could be that we did not observe substantial effects on elongation because the energetics of curved to straight transitions on the microtubule end are not rate-contributing for elongation, perhaps because these transitions occur subsequent to end binding.

The significant mutant-induced structural changes in the lattice were not accompanied by equivalent changes in the conformation of unoligomerized αβ-tubulin. The differential response to the mutation inside and outside of microtubules (Figure 7A,B) is consistent with the view that the curved conformation represents the ‘ground state’ of αβ-tubulin independent of nucleotide state (Rice et al., 2008; Buey et al., 2006; Nawrotek et al., 2011; Pecqueur et al., 2012; Ayaz et al., 2012), that the nucleotide acts across the longitudinal interface (Rice et al., 2008; Nawrotek et al., 2011), and that nucleotide-dependent interactions with the microtubule lattice are what drive αβ-tubulin conformational transitions (Rice et al., 2008; Buey et al., 2006; Nawrotek et al., 2011). Indeed, the changes we observed can largely be explained by an impaired allosteric response to GDP in the lattice. It will be interesting to discover in future work if other mutations in β-tubulin can straighten unpolymerized αβ-tubulin or modulate the allosteric response(s) to nucleotide in the lattice, and to determine if mutations in α-tubulin can yield similar effects.

In summary, we showed that buried mutations of or near β:T238 alter the allosteric response to GDP in the microtubule lattice, with dramatic consequences for catastrophe frequency and shrinking rate. By describing microtubules with identical interface composition that nevertheless undergo strikingly different polymerization dynamics, our data demonstrate that allostery in the lattice dictates functionally important aspects of microtubule polymerization dynamics (Figure 7C). Based on the central role of allostery in controlling the frequency of catastrophe and the rate of shrinking, we speculate that cooperative conformational linkage in the lattice amplifies the individual response to nucleotide. Together, the allosteric response to GDP and the intrinsic bias of αβ-tubulin toward the curved conformation elevate and separate the threshold concentrations for persistent elongation and for spontaneous nucleation (Figure 7C), and underlie the fast shrinking rate that makes catastrophe more decisive. These allosteric contributions to microtubule dynamics make microtubules more amenable to regulation by cellular factors that enhance elongation, trigger catastrophe, and promote nucleation.

Materials and methods

Protein expression and purification

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Plasmids to express TOG1, TOG2 and wild-type yeast αβ-tubulin were previously described (Johnson et al., 2011; Ayaz et al., 2012; 2014). A Bim1-GFP construct, in pHAT vector containing C-terminal EGFP-tag followed by a Strep-tag II, was a gift from Dr. Gary Brouhard. A plasmid to express the T238A mutation of Tub2p (yeast β-tubulin) was made by QuikChange (Stratagene) mutagenesis, using an expression plasmid for wild-type Tub2 as template and with primers designed according to the manufacturer’s instructions. A C-terminal FlAsH (Griffin et al., 2000) sequence was added to a Tub1 construct using polymerase chain reaction with primers designed to add the sequence WDCCPGCCK (Griffin et al., 2000). The integrity of all expression constructs was confirmed by DNA sequencing.

Wild-type or mutant yeast αβ-tubulin was purified from inducibly overexpressing strains of S. cerevisiae using Ni-affinity and ion exchange chromatography (Johnson et al., 2011; Ayaz et al., 2012; 2014) with the exception that T238A and wild-type FlAsH mutants were eluted from the Ni-affinity column with 200 mM NaCl. Prior to ion exchange chromatography, T238A Ni elution fractions were treated with Universal Nuclease (Pierce) at RT for 1 hr. Tubulin samples were stored in storage buffer (10 mM PIPES pH 6.9, 1 mM MgCl2, 1 mM EGTA) containing 20 or 50 µM GTP depending on the application. The TOG1(1-317) and TOG2(318-560) were expressed in E. coli with C-terminal His6 tags and purified using Ni-affinity, gel filtration (TOG1) and ion exchange chromatography (TOG2) (Ayaz et al., 2012; 2014). TOG domains were stored in RB100 (25 mM Tris pH 7.5, 100 mM NaCl, 1 mM MgCl2, 1 mM EGTA). Expression of Bim1-GFP was induced in the BL21(DE3) strain of E. coli with 0.5 mM IPTG for 15 hrs at 16C. Single pellets were resuspended in 50 mM Na2HPO4, 300 mM NaCl, 40 mM imidazole and sonicated for 30 min in the presence of PMSF. Lysates were clarified by centrifugation. Cleared lysate was loaded onto a His60 Superflow Column (Clontech) and the final sample was eluted in 200 mM imidazole. Elution fractions were loaded onto a 3 mL Strep-Tactin Superflow column (IBA, Germany) and eluted in RB100 with 5 mM desthiobiotin.

Yeast strains

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For expression:

JEL1 MATα leu2 trp1 ura3−52 prb1−1122 pep4−3 Δhis3::PGAL10-GAL4

For analysis of growth phenotypes:

CUY 2172 MATa TUB2::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2179 MATa tub2-T238A::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2198 MATa tub2-T238S::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2200 MATa tub2-T238V::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

For microtubule imaging:

CUY 2265 MATa TUB2::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2238 MATa tub2-T238A::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2208 MATa tub2-T238S::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

CUY 2209 MATa tub2-T238V::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52

In vivo experiments

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tub2 alleles were integrated into yeast as described previously (Reijo et al., 1994) in vivo imaging of microtubule dynamics was performed as described previously (Huang and Huffaker, 2006). Note that introduction of GFP-TUB1 into strains containing the tub2-T238A mutation lessens their dependence on benomyl for growth (Figure 6) and allowed us to image these cells in the absence of benomyl.

Time-lapse measurements of microtubule dynamics

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Flow chambers were prepared as described previously (Gell et al., 2010), with the exception that sea urchin axonemes (Waterman-Storer, 2001) were used to template yeast MT growth. Chambers were rinsed with BRB80 (80 mM PIPES pH 6.9, 1 mM MgCl2, 1 mM EGTA), followed by 10 min incubation with sea urchin axonemes. Chambers were then blocked with 1% F-127 Pluronic in BRB80 for 5 min, and washed with 1X PEM (100 mM PIPES pH 6.9, 1 mM EGTA, 1 mM MgSO4) containing 1 mM GTP. Reaction chambers were sealed with VALAP after addition of αβ-tubulin. Wild type or mutant yeast αβ-tubulin (in storage buffer containing 50 µM GTP) was taken from -80˚C, rapidly thawed, and passed through a 0.1 µm centrifugal filter at 4˚C to remove aggregates. The concentration of αβ-tubulin was measured by UV absorbance using an extinction coefficient of 115000 M-1cm-1. Protein was kept on wet ice for no more than 30 min before measuring MT dynamics. MT dynamics were imaged by differential interference contrast microscopy (DIC) using an Olympus IX81 microscope with a Plan Apo N 60x/1.42 NA objective lens and DIC prisms. Illumination at 550 nm was obtained by inserting a bandpass filter of 550/100 nm (Olympus) in the light path. Temperature was maintained at 30˚C using a WeatherStation temperature controller with enclosure fit to the microscope’s body. Micro-Manager 1.4.16 (Edelstein et al., 2010) was used to control the microscope and a Hamamatsu ORCA-Flash2.8 CMOS camera used to record the reactions. MT dynamics were recorded by taking an image every 500 ms for 1 to 2 hrs. At the end of each movie, a set of 100 out-of-focus background images was taken for background subtraction (see below). To improve signal to noise, batches of 10 raw images were averaged using ImageJ (Schneider et al., 2012) and intensity normalized before background subtraction. MT length was measured manually using a PointPicker plugin for ImageJ. Rates of MT elongation and catastrophe frequencies were determined as described previously (Walker et al., 1988).

Electron microscopy

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To prepare microtubules for electron microscopy, samples of wild-type (3-–4 µM) or β:T238A αβ-tubulin (2 µM) were prepared in 100 mM PIPES pH 6.9, 10% glycerol, 2 mM MgSO4, 0.5 mM EGTA, 1 mM GTP and incubated at 30 ˚C for 1 hr. 5 µl of the assembly reactions were spotted onto freshly glow-discharged 400 mesh grids with a carbon coated formvar support (Ted Pella), incubated for 30 s, rinsed with water, and stained with 2% aqueous uranyl acetate. Negatively stained grids were imaged at 23,000 x magnification using a Tecnai G2 Spirit electron microscope equipped with a 2Kx2K CCD camera (Gatan).

Assays for microtubule nucleation

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Microtubule sedimentation assays were performed using a range of wild-type or mutant yeast αβ-tubulin concentrations (Figure 2B, 0.25 – 4 μM) in assembly buffer (100 mM PIPES pH 6.9, 10% glycerol, 2 mM MgSO4, 0.5 mM EGTA) containing the indicated nucleotide. Samples were polymerized for 90 min at 30˚C. For SDS-PAGE analysis, microtubules were pelleted by centrifugation at 60,000 rpm (∼150000 x g) at 30˚°C for 30 min in a pre-warmed TLA-100 rotor (Beckman-Coulter), supernatant was carefully removed, and the pellet was re-suspended in an equal volume of assembly buffer such that the pellet and supernatant fractions were of equal volume. To image the products of the nucleation reactions, the reactions were cross-linked by diluting 10-fold into assembly buffer containing 1% glutaraldehyde. Cross-linking was quenched after 3 min by 5-fold dilution into assembly buffer containing 20 mM Tris pH 6.8. 150 μL of the quenched, cross-linked reactions were applied to the top of a glycerol cushion (20% glycerol in BRB80) and spun through the cushion onto poly-L-lysine coated coverslips. Coverslips were washed with BRB80, fixed with methanol and stained using FITC-DM1α (Sigma-Aldrich) for imaging by epifluorescence, as described previously (Ayaz et al., 2012).

Determining nucleotide content

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Wild type (2 or 1 µM respectively for GTP or GTPγS) or β:T238A tubulin (1 µM for either nucleotide) was mixed with 100 µM GTP or GTPγS (containing 33 nM [α-32P or γ-32P]-GTP) in assembly buffer (see above) on ice. Microtubules were assembled and harvested by centrifugation at 80,000 rpm (∼268000 x g) at 30˚C for 10 min in a pre-warmed TLA-120 rotor (Beckman-Coulter). Supernatant was carefully removed, and the pellets were gently washed 4 times with pre-warmed assembly buffer, and then resuspended with 6 M guanidine to denature the protein and release bound nucleotide. After 10-fold dilution into water, samples were loaded onto a Cellulose PEI TLC plate (Selecto Scientific) and TLC was performed, first with water followed by buffer containing 0.75 M Tris, 0.4 M LiCl, and 0.45 M HCl. The TLC plate was exposed to X-ray film after air drying. Radiolabelled mixtures of GTP/GDP and GTP/Pi were used as markers.

Analytical ultracentrifugation

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Samples for analytical ultracentrifugation (TOG1, TOG2, yeast αβ-tubulin, and mutant T238A polymerization-competent yeast αβ-tubulin) were dialyzed into final buffer conditions of RB100 (25 mM Tris pH 7.5, 1 mM MgCl2, 1 mM EGTA, 100 mM NaCl) containing 20 µM GTP. The samples shown in Figure 3 contain 0.15 µM T238A yeast αβ-tubulin and 0.015 µM, 0.03 µM, 0.15 µM, 0.3 µM, 0.6 µM, 1.2 µM, 5 µM TOG1 or 0.015 µM, 0.025 µM, 0.05 µM, 0.1 µM, 0.2 µM, 0.4 µM, 0.8 µM, 1.6 µM, and 5 µM TOG2. Samples were mixed and incubated at 4˚C for at least one hr prior to the experiment. All analytical ultracentrifugation experiments were carried out in an Optima XL-I centrifuge using an An50-Ti rotor (Beckman-Coulter). Approximately 390 µL of each sample were placed in charcoal-filled, dual-sector Epon centerpieces. Sedimentation (rotor speed: 50,000 rpm) was monitored using absorbance at 229 nm and centrifugation was conducted at 20˚°C after the centrifugation rotor and cells had equilibrated at that temperature for at least 2.5 hrs. Protein partial-specific volumes, buffer viscosities, and buffer densities were calculated using SEDNTERP (Laue et al., 1992). Data were analyzed using SEDFIT and SEDPHAT (available at http://www.analyticalultracentrifugation.com) (Schuck, 2000; Schuck et al., 2002).

Microtubule depolymerization fluorescence assays

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Wild-type and mutant yeast αβ-tubulin were labeled with 6-–10 µM ReAsH in DMSO (Life Technologies) and 1 mM TCEP, in tubulin storage buffer (see above) for 90 min at RT. To remove excess unbound dye, samples were exchanged into assembly buffer (100 mM PIPES pH 6.9, 10% glycerol, 2 mM MgSO4, 0.5 mM EGTA) with 2 mL, 7K MWCO Zeba spin desalting columns (Thermo Scientific). Labeled wild-type and mutant yeast αβ-tubulin were polymerized in the presence of 3–5 µM epothilone-B in assembly buffer (see above) with 1 mM GTP or in the presence of 500 µM GTPγS in assembly buffer (see above). The mixture was incubated for 30 min at 30 ˚°C.

Flow chambers were prepared as described above. His-Tag Antibody (1:200, Gentech) was incubated in the chamber for 10 min, followed by incubation with 1% Pluronic F-127 in BRB80 for 5 min, followed by a wash with BRB80. Pre-formed, epothilone- or GTPγS-stabilized wild-type or T238A yeast MTs were then introduced into the chamber and allowed to incubate for 10 min, followed by a wash with BRB80 to remove unbound MTs. Solutions containing a range of TOG1 (0.1-–10 µM) or TOG1(R200A) (250, 750 nM) concentrations (Figure 5B) in imaging buffer (BRB80 + 200 nM epothilone + 0.1 mg/mL BSA + antifade reagents (glucose, glucose oxidase, catalase), without the addition of β-mercaptoethanol [Gell et al., 2010]) were introduced into the chamber immediately prior to data collection. To ensure that the TOG-induced depolymerization was not an artifact of fluorescence imaging, unlabeled wild-type microtubule were assembled as described above but without ReAsH labeling and then imaged with two concentrations of TOG1 using DIC microscopy (black points in Figure 5B). MT depolymerization reactions with fluorescent microtubules were imaged by epifluoresence microscopy using an Olympus IX81 microscope with a Plan Apo N 60x/1.42 NA objective lens and Hamamatsu ORCA-Flash2.8 CMOS camera, a mercury short arc lamp, and a Texas Red filter cube (Olympus). Reactions were temperature controlled and the microscope was controlled as described above. Images of MT shrinking were recorded every 60 s for about 10 min. MT depolymerization reactions with non-fluorescent wild-type microtubules were imaged using differential interference microscopy as described above. MT length was measured manually using ImageJ (Schneider et al., 2012). Average lengths of MT’s were taken over the time course of the movie and were used to determine rate of depolymerization over an hour time span.

Dynamic assays with Bim1-GFP

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Preparation of flow chambers using sea urchin axonemes was followed as described above. Samples of wild-type, β:T238A, or C354A αβ-tubulin with 1 mM GTP or GTPγS were prepared and incubated in flow-chambers for 30–180 min and observed under DIC to evaluate presence of microtubule growth. After a given time, solutions containing desired tubulin sample and nucleotide of interest, along with 5 or 50 nM Bim1-GFP with imaging buffer described above, were flowed into the chamber. Interactions of Bim1-GFP with MTs were imaged by total internal reflection fluorescence microscopy using an Olympus IX81 microscope with a TIRF ApoN 60x/1.49 objective lens, a 491 nm 50 mW solid-state laser and Hamamatsu ORCA-Flash2.8 CMOS camera (Olympus). Reactions were temperature controlled at 30°C and the microscope was controlled as described above. Axonemes were tracked under DIC and TIRF conditions. Images of MTs were taken over several frames from 15–30 min. Bim1-GFP fluorescence intensity along microtubules and extending beyond their growing ends was obtained using the PlotProfile function in ImageJ (Schneider et al., 2012). These linescans were manually aligned to superimpose the sharp change in intensity at the very microtubule end, and aligned intensity values were averaged.

Analysis of sequence conservation

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To collect a large number of α- and β-tubulin sequences, we performed psiblast (Altschul et al., 1997) (non-redundant (nr) database, 3 iterations) on the amino acid sequences of Tub1p and Tub2p from S. cerevisiae. The top 1000 sequences from both searches were aligned using Clustal Omega (Sievers et al., 2011) and manually pruned in Jalview (Waterhouse et al., 2009) to eliminate sequences with large insertions or deletions.

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Decision letter

  1. Thomas Surrey
    Reviewing Editor; LRI, United Kingdom

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled "A mutation uncouples the tubulin conformational and GTPase cycles, revealing allosteric control of microtubule dynamics" for peer review at eLife. Your submission has been favorably evaluated by Vivek Malhotra (Senior editor), a guest Reviewing editor, and three reviewers.

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

Summary:

This manuscript describes the characterization of a T238A mutation in beta-tubulin of S. cerevisiae. In mutant cells, the microtubules are highly stabilized, and the purified mutant tubulin forms microtubules that depolymerize very slowly. The authors also determine that subsequent to polymerization, GTP is however hydrolyzed by mutant tubulin more efficiently than in microtubules polymerized from wild type tubulin. Surprisingly, EB1 decoration is not restricted to the microtubule end, suggesting that the conformational change in response to GTP hydrolysis in this mutant is different from that in normal microtubules. Similar behavior is observed for another mutation in helix 7 of beta-tubulin, consistent with prior reports (e.g. Alushin et al, 2014) that this helix is involved in the conformational changes in tubulin following hydrolysis of GTP. This is the first thorough in vitro characterization of a mutation in tubulin that uncouples GTPase cycle reactions from a conformational transition.

Essential revisions:

Although the majority of reviewers found this study very interesting and significant, some concerns were raised regarding the mechanistic insight, the way the manuscript presents its major conclusions and some technical issues of the assays as presented. These issues need to be addressed and are listed below.

Specific points of major criticism:

I) Presentation of conclusions: In its generality the novelty of the observed uncoupling of GTPase transitions and allosteric conformational changes appeared somewhat exaggerated given that drugs like taxol are well known to cause such uncoupling; this should be acknowledged and the conclusions stated more precisely. At times observations were not clearly separated from interpretations (TOG binding–conformational changes); more precise language is encouraged throughout. At times relevant literature was not acknowledged: high GTP content in yeast tubulin: Dougherty et al, Biochemistry 37, 1998, p. 10861; importance of the H7 helix for microtubule stability: Amos, Org Biomol Chem 2, 2004, p. 2153.

II) Technical quality/support of conclusions by the data: Several technical concerns have been identified that require additional experiments:

1) Figure 3 – TOG binding curves. The quality of the assay was not considered sufficient (large errors, saturation not reached) to support the strong statement that the mutant does not change TOG binding, leading to the interpretation that it does not change the conformation of unpolymerised tubulin. Additional data points need to be collected to bring the errors down to support such a strong statement convincingly.

2) Figure 4 – Bim1 end tracking: More example of images or kymographs of Bim1 end tracking on wildtype and mutant microtubules should be shown to demonstrate how 'typical' behavior looks like under the conditions used. Bim1 binding to mutant microtubules is only shown for one Bim1 concentration. Therefore, it remains unclear if simply the overall affinity of Bim1 for microtubule binding is increased or if indeed the relative accumulation at microtubule ends compared to the lattice is reduced/is now absent (as the authors conclude). Experiments with different Bim1 concentrations are needed to distinguish between these two possibilities (effect on overall affinity versus end binding selectivity).

3) Figure 5 – depolymerization of mutant microtubules by TOG: Technical concerns regarding the assay conditions have been raised. It was considered possible that the strong depolymerization activity of TOG could maybe be a consequence of artefacts due to singlet oxygen generation by ReAsH in the absence of reducing agents/strong oxygen scavenging system. The possibility of such effects should be excluded experimentally (changing overall fluorescence excitation conditions, adding oxygen scavengers, or changing fluorophore).

4) Figure 1c – dependence of growth velocities on tubulin concentration: the data for mutant versus wildtype microtubules have been collected in very different tubulin concentration regimes (due to the strong nucleation of mutant tubulin). So the strong conclusion that this dependence is identical for both types of microtubules relies on how reliably the individual curves can be extrapolated into the other regime/how big the errors of the fits are. These errors should be examined in more detail. Ideally, an attempt should be made to collect data points for wildtype microtubules close to its critical concentration using a recently published method (Wieczorek et al., Nat Cell Biol 17, 2015, p. 907).

5) One reviewer felt strongly that simple negative stain EM should be performed to demonstrate that the microtubule structure appears normal for mutant microtubules. Abnormalities not visible by fluorescence microscopy can then be excluded. This would set the standard for other future studies with mutant tubulins.

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

Thank you for resubmitting your work entitled "A mutation uncouples the tubulin conformational and GTPase cycles, revealing allosteric control of microtubule dynamics" for further consideration at eLife. Your revised article has been favorably evaluated by Vivek Malhotra (Senior editor) and a guest Reviewing editor. The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance, as outlined below.

Most major points of criticism have been addressed satisfactorily by additional experiments and the presentation has been improved. At several instances however the clarity of presentation would still profit from further improvement and one essential control experiment is missing:

I) Missing control experiment:

Figure 4: The new data clearly strengthen the manuscript. The schemes and the language used here continue to suggest that the GTPgS lattice is identical to the mutant lattice. However, the new data demonstrate that there is a difference between the mutant GDP microtubule body and the GTP end. This may well indicate that there may also be a difference between the mutant GDP body and the GTPgS body (GTPgS being potentially more similar to the end than to the mutant body). A control experiment with 5 nM Bim1 on GTPgS microtubules is missing. The result of this experiment might require the authors to modify the scheme and adopt a more differentiated view about the microtubule lattice conformations of wt, mutant and GTPgS microtubules.

II) Presentation:

1) Abstract and text: The language of the authors continues to imply complete uncoupling between GTPase reaction and conformational changes, although the new Bim1 data in Figure 4 demonstrate that there is clearly a conformational change in the mutant in response to GTPase, however at a reduced level as compared to wild type. It is desirable to adjust language to reflect this point. For the same reason, the claim made in the Abstract that "post-GTPase conformational changes are not strictly required for catastrophe" does not appear to be supported by the new data and it is recommended to adjust language.

2) Yeast strains: The genotypes of the strains used for the tubulin productions and live cell microscopy experiments should be stated (Methods, Legends and/or Table).

3) Figure 1: The number of observed events used for the quantifications does not appear to be stated. Please check throughout if all numbers relevant for statistical analysis are provided.

4) Figure 1A, Figure 6C: Please add information to the figures to orient the reader where the inside/outside, plus/minus end of the microtubule is.

5) Figure 1D: Please provide clearer examples of kymographs. Contrast is very low.

6) Figure 2: The tubulin concentrations used for the data presented in Figure C,D does not seem to be provided. Please put this concentration into context of the concentrations used in A and the discussion about the oligomers that might lead to the measurement of an increased GTP level.

7) Figure 4B: Please explain how alignment and averaging of the Bim1 intensity profiles was performed. Please clarify if error bars are shown (the GTPgS graph displays some fuzzy haze which might be bars) and adjust the display so that the data can be seen clearly. Please check the manuscript throughout and state which errors are shown (s.d. or s.e.m.).

8) Figure 5: "Similar results were obtained using MTs stabilized with GTPgS (not shown)." Please show the existing data to support the claim.

9) The conclusions drawn here should be discussed in light of a recent publication from the Nogales lab (Cell 162, 849, 2015) which contains relevant information about the structure of microtubules in the absence and presence of an end binding protein and different nucleotides. Some of the interpretations might need refinement.

https://doi.org/10.7554/eLife.10113.016

Author response

Although the majority of reviewers found this study very interesting and significant, some concerns were raised regarding the mechanistic insight, the way the manuscript presents its major conclusions and some technical issues of the assays as presented. These issues need to be addressed and are listed below.

We appreciate that the reviewers found our work to be interesting and significant. In brief we have tuned and tightened the language throughout, and we have added new data and/or panels to Figures 1, 3, 4, and 5. The revised manuscript is stronger and clearer, and substantively addresses all of the concerns articulated in the decision letter.

Specific points of major criticism: I) Presentation of conclusions: In its generality the novelty of the observed uncoupling of GTPase transitions and allosteric conformational changes appeared somewhat exaggerated given that drugs like taxol are well known to cause such uncoupling; this should be acknowledged and the conclusions stated more precisely. At times observations were not clearly separated from interpretations (TOG binding–conformational changes); more precise language is encouraged throughout. At times relevant literature was not acknowledged: high GTP content in yeast tubulin: Dougherty et al, Biochemistry 37, 1998, p. 10861; importance of the H7 helix for microtubule stability: Amos, Org Biomol Chem 2, 2004, p. 2153.

We completely agree that small molecules such as taxol are already known to uncouple the GTPase and conformational cycles, and we did not intend to appear to lay claim to having discovered this for the first time. We have changed the Abstract, Introduction, Results, and Discussion sections to be clearer and more precise about what is different and novel in our work.

We have also added the suggested citations and also tightened the language throughout.

II) Technical quality/support of conclusions by the data: Several technical concerns have been identified that require additional experiments: 1) Figure 3 – TOG binding curves. The quality of the assay was not considered sufficient (large errors, saturation not reached) to support the strong statement that the mutant does not change TOG binding, leading to the interpretation that it does not change the conformation of unpolymerised tubulin. Additional data points need to be collected to bring the errors down to support such a strong statement convincingly.

We have collected additional data points for the TOG2 titration, which slightly increased the fitted affinity and slightly decreased its error range. Lack of saturation was not a factor. A simple thought experiment will illuminate why these data are sufficient to support an argument that the mutation does not lead to substantial change in αβ−tubulin curvature. Based on our measurements, TOG domains bind approximately 2-fold tighter to β:T238A αβ-tubulin than they do to wild-type. A 2-fold difference in binding constant is equivalent to about 0.7 kT of binding energy – this is a similar order of magnitude as one hydrogen bond, and therefore hard to square with a substantial conformational rearrangement in the mutant. We can also estimate the order of magnitude change in binding affinity we might expect for curved vs straight αβ-tubulin. ~100 nM affinity corresponds to about 16 kT of total binding energy. TOG domains contact both α- and β- tubulin when binding to curved αβ-tubulin, but in straight αβ-tubulin TOG domains cannot maintain both contacts simultaneously. Making a simplifying assumption that about half of the binding energy (8 kT) will be lost for the straight conformation would correspond to ~3000-fold weaker binding, an estimate that is consistent with the fact that we have not observed significant binding of TOGs to microtubules or detectable binding to αβ-tubulin by a ‘half-site’ TOG1 mutant (R200A). Thus, the fitted affinities and their associated errors are not consistent with the idea that there is a substantial mutation-induced change in TOG binding affinity, and this in turn supports our claim that the conformation of the mutant cannot be very different than that of the wild-type. We feel that this is especially so given that we readily detected a change in Bim1 binding affinity for the body of mutant microtubules, and that we also observed a nearly two order of magnitude change in depolymerization rate, all with only small detectable changes in the apparent biochemistry of elongation.

In the original submission, we had tried to be careful throughout the manuscript to state that the mutant did not detectably change the conformation of unpolymerized αβ-tubulin. In the revised manuscript we have maintained the use of qualifiers to indicate the lack of complete certainty (which would likely require structures of mutant and wild-type αβ- tubulin, and more) while also re-working the discussion surrounding these data to include some of the arguments from the prior paragraph. We appreciate the chance to clarify these issues.

2) Figure 4 – Bim1 end tracking: More example of images or kymographs of Bim1 end tracking on wildtype and mutant microtubules should be shown to demonstrate how 'typical' behavior looks like under the conditions used.

We have now included more images of Bim1 end tracking on wild-type and mutant microtubules (4 for each condition as opposed to 1 previously; we also added two new conditions, see below).

Bim1 binding to mutant microtubules is only shown for one Bim1 concentration. Therefore, it remains unclear if simply the overall affinity of Bim1 for microtubule binding is increased or if indeed the relative accumulation at microtubule ends compared to the lattice is reduced/is now absent (as the authors conclude). Experiments with different Bim1 concentrations are needed to distinguish between these two possibilities (effect on overall affinity versus end binding selectivity).

This comment raises interesting ideas about the microtubule cap. We have performed additional experiments in response to it. In our initial submission the Bim1 binding clearly showed that the body of mutant microtubules presents higher-affinity Bim1 binding sites than does the body of wild-type microtubules. This comment asks us to determine if the same is true in the cap region. New experiments using 5 nM Bim1 (10-fold lower concentration than in the original submission) show that (i) the Bim1 intensity in the cap is comparable for mutant and wild-type microtubules, suggesting that wild-type and mutant caps present Bim1 binding sites of comparable affinity, (ii) the Bim1 intensity on the body of mutant microtubules is significantly higher than on the body of wild-type microtubules, consistent with our proposal that αβ-tubulin adopts a more expanded conformation in β:T238A microtubules, and (iii) we see ‘comets’ on the end of mutant microtubules, indicating that Bim1 binds more tightly to the ends of mutant microtubules than to the body.

Our bottom-line conclusion that β:T238A αβ-tubulin adopts a more expanded, GTP-like conformation in a GDP lattice remains unchanged. However, by showing that Bim1 binds to mutant microtubule caps more tightly than to mutant microtubule bodies, the new data now make it very clear that the mutation does not completely override the nucleotide effect. We have modified the Discussion to reflect these new findings, including substituting ‘suppress’ for more absolute words like ‘eliminate’. In our view these new data and associated discussion strengthen the manuscript, and we are grateful for the push to do these experiments.

3) Figure 5 – depolymerization of mutant microtubules by TOG: Technical concerns regarding the assay conditions have been raised. It was considered possible that the strong depolymerization activity of TOG could maybe be a consequence of artefacts due to singlet oxygen generation by ReAsH in the absence of reducing agents/strong oxygen scavenging system. The possibility of such effects should be excluded experimentally (changing overall fluorescence excitation conditions, adding oxygen scavengers, or changing fluorophore).

The assays reported in our initial submission did in fact include a strong oxygen scavenging system (Glucose oxidase + catalase), but we can see how this might have been overlooked because we lumped them into the phrase ‘antifade reagents’. We now mention these reagents explicitly, and we also performed two kinds of control experiments to rule out the possibility of a non-specific origin of the effects we observed. First, we monitored the rate of TOG-induced shrinking using differential interference contrast microscopy at two different TOG concentrations and without any fluorescent labels. These experiments gave microtubule shrinking rates consistent with what we initially reported. Second, using the ReAsH labeled microtubules, we performed experiments using a TOG domain containing a mutation that greatly reduces tubulinbinding affinity. These experiments revealed greatly reduced rates of TOG-induced shrinking. These new data have been added to Figure 5B. Together, they rule out the possibility that the TOG-induced depolymerization was an artifact.

4) Figure 1c – dependence of growth velocities on tubulin concentration: the data for mutant versus wildtype microtubules have been collected in very different tubulin concentration regimes (due to the strong nucleation of mutant tubulin). So the strong conclusion that this dependence is identical for both types of microtubules relies on how reliably the individual curves can be extrapolated into the other regime/how big the errors of the fits are. These errors should be examined in more detail. Ideally, an attempt should be made to collect data points for wildtype microtubules close to its critical concentration using a recently published method (Wieczorek et al., Nat Cell Biol 17, 2015, p. 907).

We had examined the fitting errors but somehow managed to not include this in the manuscript. We have now corrected this oversight by including the fitting errors and by discussing them when we are comparing the slopes and intercepts of the two fitted lines. To minimize the ‘distance’ of extrapolation, we also obtained an additional data point for wild-type αβ-tubulin using the Wieczorek/Brouhard method. We apologize for not including and discussing fitting errors in our initial submission.

5) One reviewer felt strongly that simple negative stain EM should be performed to demonstrate that the microtubule structure appears normal for mutant microtubules. Abnormalities not visible by fluorescence microscopy can then be excluded. This would set the standard for other future studies with mutant tubulins.

We have now included negative stain EM images showing that wild-type and β:T238A microtubules display similar structure.

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

I) Missing control experiment: Figure 4: The new data clearly strengthen the manuscript. The schemes and the language used here continue to suggest that the GTPgS lattice is identical to the mutant lattice. However, the new data demonstrate that there is a difference between the mutant GDP microtubule body and the GTP end. This may well indicate that there may also be a difference between the mutant GDP body and the GTPgS body (GTPgS being potentially more similar to the end than to the mutant body). A control experiment with 5 nM Bim1 on GTPgS microtubules is missing. The result of this experiment might require the authors to modify the scheme and adopt a more differentiated view about the microtubule lattice conformations of wt, mutant and GTPgS microtubules.

We had chosen to focus on the differential conformational response to GDP, so we did not perceive the 5 nM Bim1 on GTPgS microtubules as a missing control because GTPgS microtubules have uniform nucleotide state. We now show those data in the revised Figure 4, and they document the expected result that Bim1 coats the body of GTPgS microtubules with similar intensity as it does the tip. To avoid conveying an impression that the mutant lattice is identical to a GTP/GTPgS lattice, we have made changes to the language throughout the paper (see also our response to the next point).

II) Presentation: 1) Abstract and text: The language of the authors continues to imply complete uncoupling between GTPase reaction and conformational changes, although the new Bim1 data in Figure 4 demonstrate that there is clearly a conformational change in the mutant in response to GTPase, however at a reduced level as compared to wild type. It is desirable to adjust language to reflect this point. For the same reason, the claim made in the Abstract that "post-GTPase conformational changes are not strictly required for catastrophe" does not appear to be supported by the new data and it is recommended to adjust language.

In the revised text we submitted we had inserted a number of qualifying statements in an effort to not portray the ‘uncoupling’ in a black and white manner. We have now done even more of that, adopting phrasing that is closer to the biochemical finding (e.g. stronger Bim1 binding, GTP-lattice-like character) in order to avoid too many statements about specific conformations (expanded/compacted). We also modified the ‘strictly required’ sentence in the Abstract, which we should have done the first time around. Thanks for catching that, and apologies for missing it.

2) Yeast strains: The genotypes of the strains used for the tubulin productions and live cell microscopy experiments should be stated (Methods, Legends and/or Table).

Strain genotypes have been added as a separate section in the Methods.

3) Figure 1: The number of observed events used for the quantifications does not appear to be stated. Please check throughout if all numbers relevant for statistical analysis are provided.

Done in Figure 1 and throughout. As part of this checking we changed some panels in Figure 1 to show s.e.m instead of s.d.

4) Figure 1A, Figure 6C: Please add information to the figures to orient the reader where the inside/outside, plus/minus end of the microtubule is.

We have explained the viewing direction in the legends of the figures where we show views of the structures.

5) Figure 1D: Please provide clearer examples of kymographs. Contrast is very low.

Done.

6) Figure 2: The tubulin concentrations used for the data presented in Figure C,D does not seem to be provided. Please put this concentration into context of the concentrations used in A and the discussion about the oligomers that might lead to the measurement of an increased GTP level.

We have listed the concentrations in the Methods and in the figure legend and in the text, where we also note that they are within the range shown in panel A. We did not add additional discussion, in part because panels A, B, and C each used different centrifugation settings. We did adjust the language we used to discuss possible origins of the increased GTP level.

7) Figure 4B: Please explain how alignment and averaging of the Bim1 intensity profiles was performed. Please clarify if error bars are shown (the GTPgS graph displays some fuzzy haze which might be bars) and adjust the display so that the data can be seen clearly. Please check the manuscript throughout and state which errors are shown (s.d. or s.e.m.).

We have added to the methods section to better explain what we did. We agree that the error bars were not clearly visible and we have altered the formatting to improve the presentation of the data. Legends now state if error bars represent s.d or s.e.m.

8) Figure 5: "Similar results were obtained using MTs stabilized with GTPgS (not shown)." Please show the existing data to support the claim.

We have added these data points to Figure 5.

9) The conclusions drawn here should be discussed in light of a recent publication from the Nogales lab (Cell 162, 849, 2015) which contains relevant information about the structure of microtubules in the absence and presence of an end binding protein and different nucleotides. Some of the interpretations might need refinement.

Two factors make it difficult to draw direct comparisons between our results and the three lattice structures recently described by Eva Nogales and colleagues. First, we don’t yet have high-resolution structures for wild-type or mutant yeast microtubules, so we can’t know exactly which conformations are involved. Second, yeast microtubules assemble readily using GTPgS but less so with GMPCPP, whereas the opposite is true for vertebrate microtubules – GTPgS may therefore be promoting different states for the two kinds of microtubules. These factors limit our ability to make detailed structural interpretations, and as mentioned above we have made changes to our language to try to make this clear. Nevertheless, the different Bim1 binding properties of wild-type and mutant microtubules provide strong support for the idea that the mutant retains significant GTP-lattice-like character in a GDP lattice. Encouraging more explanation around these ideas was a good suggestion, and we have added a few sentences along the lines of the above to the end of the Discussion.

https://doi.org/10.7554/eLife.10113.017

Article and author information

Author details

  1. Elisabeth A Geyer

    Departments of Biophysics and Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    EAG, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Alexander Burns

    Departments of Biophysics and Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    AB, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  3. Beth A Lalonde

    Department of Molecular Biology and Genetics, Cornell University, Ithaca, United States
    Contribution
    BAL, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  4. Xuecheng Ye

    Departments of Biophysics and Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    XY, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  5. Felipe-Andres Piedra

    Departments of Biophysics and Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    FAP, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  6. Tim C Huffaker

    Department of Molecular Biology and Genetics, Cornell University, Ithaca, United States
    Contribution
    TCH, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  7. Luke M Rice

    Departments of Biophysics and Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    LMR, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    Luke.Rice@UTSouthwestern.edu
    Competing interests
    The authors declare that no competing interests exist.

Funding

National Institute of General Medical Sciences (GM-098543)

  • Luke M Rice

National Science Foundation (MCB-1054947)

  • Luke M Rice

National Science Foundation (2014177758)

  • Elisabeth A Geyer

National Institute of General Medical Sciences (T32 GM008297)

  • Elisabeth A Geyer
  • Felipe-Andres Piedra

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

Acknowledgements

We thank C. Brautigam in the UT Southwestern Macromolecular Biophysics Resource for help and advice. S Bechstedt in G Brouhard’s lab performed pilot experiments that pointed us to the TOG-induced depolymerization assay. Electron microscopy was performed in the UT Southwestern Electron Microscopy Core Facility. H Yu, X Zhang, B Russ, G Brouhard, J Kollman, and C Asbury gave insightful comments on the manuscript. LMR is the Thomas O Hicks Scholar in Medical Research. FAP and EAG. were supported by NIH T32 GM008297, and EAG. was supported by an NSF Graduate Research Fellowship, Grant No. 2014177758. This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. 2014177758. Any opinion, findings, and conclusions or recommendations expressed in this material are those of the authors(s) and do not necessarily reflect the views of the National Science Foundation. A. B. was a UT Southwestern Medical Center/UT Dallas Green Fellow. This work was supported by GM-098543 (from the NIH), and MCB-1054947 (from the NSF).

Reviewing Editor

  1. Thomas Surrey, LRI, United Kingdom

Publication history

  1. Received: July 15, 2015
  2. Accepted: October 6, 2015
  3. Accepted Manuscript published: October 6, 2015 (version 1)
  4. Version of Record published: December 9, 2015 (version 2)

Copyright

© 2015, Geyer 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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