Introduction

Microtubules (MTs) are dynamic polymers of αβ-tubulin (tubulin) that mediate essential cellular functions such as cell division, transport, and motility (Janke and Magiera, 2020). MT dynamics are regulated by multiple MT-associated proteins (MAPs) (Akhmanova and Steinmetz, 2015; Brouhard and Rice, 2018; Goodson and Jonasson, 2018; Gudimchuk and McIntosh, 2021). Defining and quantifying the biochemical mechanisms underlying MT dynamics and regulation is an ongoing challenge for the field.

Stu2/XMAP215-family proteins are conserved ‘polymerases’ (Gard and Kirschner, 1987; Ohkura et al., 1988) that catalyze MT growth using αβ-tubulin-binding TOG (Tumor Overexpressed Gene) domains (Al-Bassam et al., 2007; Ayaz et al., 2012; Brouhard et al., 2008)(Fig. 1A). All family members contain multiple, flexibly-linked TOG domains, but the domain organization and oligomerization state of these polymerases can vary across taxa (Al-Bassam and Chang, 2011; Slep, 2009)(Fig. 1B): fungal polymerases such as Stu2 (S. cerevisiae) and Alp14 (S. pombe) are homodimers containing two different TOG domains per monomer whereas metazoan polymerases like XMAP215 (X. laevis) and chTOG (H. sapiens) are monomers containing 5 (or possibly 6) different TOG domains. Despite years of study from multiple groups, a quantitative biochemical understanding of how these polymerases work has not yet been defined. Currently, two main classes of mechanism for polymerase activity have been proposed. Our group advanced a ‘concentrating reactants’ model in which catalytic polymerase activity results from linked TOG domains at the MT end recruiting unpolymerized tubulin, thereby increasing the effective concentration of tubulin and consequently the rate of tubulin:MT interactions(Ayaz et al., 2014; Ayaz et al., 2012; Geyer et al., 2018). Another group proposed a ‘polarized unfurling’ model (Nithianantham et al., 2018) in which polymerase activity results from a multi-step, multi-tubulin delivery process that postulates specific TOG:TOG contacts and distinct roles for different TOG domains; this model is based on structures of TOG:tubulin complexes but does not obviously explain why polymerase activity is catalytic. These two models represent very different views about how the polymerase might work and have different implications for thinking about TOG-based polymerase mechanism across taxa. Nevertheless, they both share a common limitation: neither quantitatively links polymerase activity to the number and tubulin-binding affinity/kinetics of TOG domains.

Figure 1.

With the goal of providing a deeper and more general understanding of Stu2/XMAP215-family polymerase mechanism and whether it is conserved between homodimeric and monomeric family members, we sought to improve over current models by defining a quantitative relationship between the growth-promoting activity of a polymerase and the number and tubulin-binding properties of its TOG domains. We were inspired by prior work on Ena/VASP, an otherwise unrelated actin polymerase that appears to share a similar functional design with Stu2/XMAP215 polymerases: both polymerases contain multiple flexibly linked domains (Figure 1C) — TOGs in Stu2/XMAP215 and GAB/EVH2 in Ena/VASP — that preferentially interact with unpolymerized tubulin and actin, respectively (Ayaz et al., 2014; Ayaz et al., 2012; Ferron et al., 2007; Walders-Harbeck et al., 2002). In contrast to the MT polymerases, however, for the actin polymerase Ena/VASP there is a quantitative biochemical model that explains how its polymerase activity results from the number and actin-binding kinetics of its GAB domains (Fig. 1D) (Breitsprecher et al., 2011). Using Stu2 as a model, we tested whether the biochemical model for an actin polymerase could also be applied to explain the activity of MT polymerases.

The biochemical model (Breitsprecher et al., 2011) essentially views the polymerase as analogous to an enzyme that acts on the polymer end, using unpolymerized subunits as its substrate. We first showed that the growth-promoting activity of Stu2 displayed two characteristics required by the biochemical model (Breitsprecher et al., 2011): (i) a fixed concentration of Stu2 stimulated MT elongation rate with hyperbolic dependence on tubulin concentration, consistent with enzyme-like action, and (ii) the amount of Stu2 on the MT end did not vary detectably with tubulin concentration, so variation in polymerase-stimulation of MT growth reflects something other than differential accumulation of the polymerase on the MT end. Finally, we directly measured the binding affinity and kinetics for TOG:tubulin interactions, revealing tighter than expected binding that is consistent with the measured dependence of polymerase activity on tubulin concentration.

The binding measurements and polymerase activity can be unified within the biochemical model, providing a quantitative understanding of how MT polymerase activity results from the number and tubulin binding kinetics of TOG domains. The results indicate that Stu2 operates with very high efficiency: the rate-limiting step in the mechanism is unpolymerized tubulin binding to a TOG domain, and once a TOG binds to an unpolymerized tubulin, that tubulin nearly always becomes incorporated into the MT. Nothing in the model requires a specific oligomerization state; given earlier data showing comparable activity for monomeric and dimeric “two TOG” variants of Stu2 (Geyer et al., 2018), the enzyme-like biochemical model should be broadly applicable to polymerases across taxa. Thus, Stu2/XMAP215-family polymerases function like tubulin-shuttling antennas that accelerate the arrival of unpolymerized tubulin to the MT end.

This kind of mechanism is more consistent with the ‘concentrating reactants’ mechanism we proposed previously (Ayaz et al., 2014). The biochemical model does not specify the mechanism of polymerase end localization, but there, too, Stu2/XMAP215 and Ena/VASP polymerases appear to share a common feature: both polymerases only localize to the growing end in the presence of unpolymerized subunits (Geyer et al., 2018; Hansen and Mullins, 2010) (in other words, end localization for both polymerases cannot be separated from polymerase activity). The similarity in mechanism between these two otherwise unrelated cytoskeletal polymerases provides an interesting example of convergent evolution. The common design principle likely indicates that localizing flexibly-linked subunit-binding domains to the growing polymer end is sufficient to generate polymerase activity.

Results

Does a similar modular organization in MT and actin polymerases reflect a common biochemical mechanism?

In seeking to define a biochemical mechanism to explain how Stu2/XMAP215-family MT polymerase activity results from the number and tubulin-binding properties of TOG domains (Fig. 1B), we were intrigued by apparent similarities between Stu2 and the actin polymerase Ena/VASP (Fig. 1C), for which a biochemical mechanism has already been defined (Fig. 1D) (Breitsprecher et al., 2011). Both Stu2 and Ena/VASP are oligomeric polymerases that contain multiple flexibly-linked domains (TOGs for Stu2/XMAP215 and GAB for Ena/VASP) that bind preferentially to unpolymerized tubulin and actin, respectively. We test here whether the model that quantitatively describes how Eva/VASP polymerase activity results from the number and biochemical properties of its actin-binding domains (Figure 1D) can also be used to provide a molecular understanding of Stu2 MT polymerase activity.

The biochemical model effectively considers the polymerase as analogous to an enzyme that acts on the polymer end and that uses unpolymerized subunits as its substrate. The biochemical model predicts a Michaelis-Menten-like (hyperbolic) dependence of polymerase-stimulated growth rate on subunit concentration, and it directly relates characteristics of polymerase activity - the apparent maximal stimulation of growth rate (V_max) and the concentration at which half-maximal stimulation is achieved (K_M) - to the number of TOG domains (N) and the rate constants for the underlying biochemical reactions: subunits associating/dissociating from a binding domain (k_on, k_off, K_D) or for being transferred from the binding domain to the growing end of the polymer (k_t) (Figure 1C). Prior in vitro measurements of Stu2/XMAP215-family polymerase activity (Brouhard et al., 2008; Cook et al., 2019; Farmer et al., 2021; Geyer et al., 2018; Podolski et al., 2014; Roostalu et al., 2015; Thawani et al., 2018; Widlund et al., 2011) have generally varied polymerase concentration at a fixed concentration of tubulin. The model being explored here considers tubulin as the substrate and consequently is most easily tested at a fixed concentration of polymerase using variable tubulin concentration. Thus, we first obtained new measurements to determine whether Stu2-stimulated growth at different tubulin concentrations showed enzyme-like characteristics, as required by the biochemical model.

Polymerase activity at variable tubulin concentrations follows Michaelis-Menten-like behavior, consistent with the biochemical model

We measured in vitro MT growth rates using a constant, roughly half-saturating amount of Stu2 (100 nM) at concentrations of yeast tubulin ranging from 0.4 to 1.4 µM. In addition to wild-type Stu2 (hereafter denoted Stu2(TOG1-TOG2) to indicate the identity of the two TOG domains per monomer), and with the goal of facilitating model fitting, we also measured two additional variants (Figure 2A): Stu2(TOG2-TOG2) has the TOG1 domain replaced by a second TOG2 domain, which simplifies the biochemistry by only having one type of TOG:tubulin interaction; Stu2(TOG1*-TOG2) has one TOG domain (TOG1) ‘inactivated’ (TOG1* denotes a variant unable to bind tubulin, achieved here using the R200A mutation (Ayaz et al., 2012)) to vary the number of active TOGs in the polymerase.

Figure 2.

Stu2 increased microtubule growth rates (Fig. 2B,C), and the Stu2-mediated growth rate (Fig. 2C) followed Michaelis-Menten-like (hyperbolic) dependence on tubulin concentration (Figure 2D). The maximal activity of Stu2(TOG2-TOG2) was comparable to that of Stu2(TOG1-TOG2) (both constructs contain 4 active TOG domains) while the maximal activity of Stu2(TOG1*-TOG2) – a construct that only contains 2 active TOG domains - was roughly half that of the unmutated constructs (Figure 2C,D; corresponding V_max and K_M values for the three constructs are summarized in Figure 2D, box). These results are consistent with prior work on these constructs (Geyer et al., 2018). That all constructs showed enzyme-like dependence on tubulin concentration means that the data may be amenable to analysis using enzyme-like biochemical model. Hereafter, experiments used Stu2(TOG2-TOG2) and Stu2(TOG1*-TOG2) constructs to avoid having to account for different TOG:tubulin affinities in the model.

The amount of Stu2 on the MT end is independent of MT growth rate

To analyze the Stu2 measurements in the context of the biochemical model requires that the amount of Stu2 on the microtubule end does not vary with tubulin concentration. We therefore quantified the amount of Stu2 on the MT end at low (0.6 µM) and high (1.4 µM) concentrations of αβ-tubulin, using TIRF to visualize GFP-tagged Stu2 on the end of growing, unlabeled MTs. End-resident Stu2 was quantified by measuring the fluorescence intensity of the Stu2-GFP comet (Figure 3A). The measured intensity did not differ detectably between low and high tubulin concentrations (Figure 3B), indicating that the amount of Stu2 on the MT end does not depend on the concentration of αβ-tubulin (or, equivalently, on how fast the microtubule is growing).

Figure 3.

The measurements reported thus far indicate that Stu2 polymerase activity shows enzyme-like characteristics that make it amenable to analysis using the biochemical model initially developed for Ena/VASP. Specifically, Stu2-stimulated MT growth depends hyperbolically on tubulin concentration (is Michaelis-Menten-like), and the amount of Stu2 on the MT end does not depend on tubulin concentration. In the biochemical model (Fig. 2D), the dependence of polymerase activity on tubulin concentration is given by the apparent K_M, which is defined by K_M = K_D + k_t/k_on. Accordingly, the concentration-dependence of polymerase activity could be dictated by the affinity of TOG:tubulin interactions (“affinity limited”: K_M ≈ K_D and k_t/k_on negligeable), by the rate constants for TOG:tubulin association (k_on) and for transfer of a TOG-bound tubulin to the polymer end (k_t) (“kinetically limited”: K_M ≈ k_t/k_on and K_M negligeable), or by a combination of both. To determine which regime is operative requires knowledge about the affinity and kinetics of TOG:tubulin interactions.

Tubulin binds rapidly and tightly to TOG2 (10 nM affinity) under the conditions of the polymerase assay

We previously measured the tubulin-binding affinity of Stu2-TOG1 (70 nM) and Stu2-TOG2 (160 nM) domains (Ayaz et al., 2014), but those equilibrium experiments did not provide information about the rates of TOG:tubulin association and dissociation. Moreover, those earlier measurements may not faithfully reflect the interactions present during the polymerase assay because they were made using different conditions of pH and ionic strength. We used Bio-layer Interferometry (BLI) to determine the affinity and kinetics of TOG2:tubulin interactions under buffer conditions close to those used for measuring polymerase activity (see Methods). Tubulin was site-specifically biotinylated on the C-terminus of β-tubulin using sortase-mediated protein splicing (Popp et al., 2007)(Fig. 4A) and then attached to streptavidin-coated chips. BLI measurements using wild-type TOG2 yielded a dose-dependent, saturable binding signal (Fig. 4B,C) and control measurements using TOG2* did not show evidence of binding (Figure 4C). Fitting the concentration-dependent amplitudes to a one-site binding model yielded an affinity of 8.9 nM for TOG2:tubulin binding (Figure 4C). Globally fitting single exponential curves to the association and dissociation phases yielded rate constants k_on (3.2 × 10⁶ M^-1s^-1) and k_off (3 × 10⁻² s^-1) with good fits (global R² = 0.97) (Figure 4B, box). Calculating the dissociation constant K_D from the measured rate constants (K_D = k_off/k_on) instead of from the amplitudes yielded a value of 9.6 nM, in good agreement with the amplitude-based determination. The ∼10 nM TOG2:tubulin binding affinity is 16-fold stronger than the 160 nM affinity measured previously at higher ionic strength (Ayaz et al., 2014), and negligeable compared to the apparent K_M for polymerase activity. Thus, the TOG domains are effectively saturated with tubulin in the concentration range we measured, and consequently the polymerase must be kinetically limited by either the rate of TOG:tubulin binding or by the rate of TOG-mediated transfer to tubulin to the microtubule end.

Figure 4.

Figure 5.

Unifying measurements of polymerase activity and TOG:tubulin interaction kinetics

Are the biochemical measurements of TOG:tubulin interactions consistent with the polymerase measurements? The biochemical model provides a way to address this question because it provides a quantitative link between the enzyme-like characteristics (K_M, V_max) of polymerase activity and the tubulin-binding and transfer reactions carried out by the TOG domains. For example, the concentration-dependence of polymerase activity is dictated by the apparent K_M, which is defined by K_M = K_D + k_t/k_on (Fig. 2D); in this expression k_t is the only unmeasured quantity, so we can solve for its value to obtain k_t = 5.04 s^-1. This transfer rate of ∼5 s^-1 for each TOG domain is close to the ∼10 s^_-1 rate at which ‘free’ (not bound to a TOG) tubulins incorporate into growing microtubules in our assay (Fig. 2C). Similarly, the maximal polymerase activity is given by V_max, which is defined by V_max = k_t*N; having measured V_max and derived k_t, we can calculate the number of TOG domains to obtain N=12 total TOG domains for Stu2(TOG2-TOG2) and N=8 total TOG domains for Stu2(TOG1*-TOG2). Since each Stu2(TOG2-TOG2) dimer contains 4 binding-competent TOGs, N=12 indicates that approximately 3 TOG-TOG2 polymerases reside at the growing microtubule end in our assay. A similar calculation for Stu2(TOG1*-TOG2), which has 2 binding-competent TOGs, yields approximately 4 polymerases at the growing microtubule end. The estimate of 3-4 polymerases on the microtubule end at ∼half-saturating concentrations of Stu2 is consistent with our earlier measurement of ∼6 Stu2(TOG1-TOG2) molecules on the microtubule end under saturating conditions (Geyer et al., 2018). Thus, the measured enzyme-like characteristics of polymerase activity are consistent with and explained by the TOG:tubulin binding kinetics.

Discussion

The experiments and analyses described here provide a quantitative biochemical mechanism that explains the enzyme-like MT polymerase activity of Stu2 in terms of the number of TOG domains and their rate-constants for binding tubulin. High-affinity (∼10 nM) TOG:tubulin interactions mean that once a TOG:tubulin complex forms, the TOG-bound tubulin will nearly always be delivered to the MT end (the 5 s^-1 rate at which each TOG-bound tubulin is transferred to the MT end is >100-fold faster than the 3 × 10⁻² s^-1 rate at which tubulin dissociates from a TOG). The polymerase therefore operates with very high delivery efficiency. The high affinity of TOG:tubulin interactions, in particular that K_D << K_M, also means that Stu2 is kinetically limited.

Our measurements indicate that the rate of tubulin binding to TOG is most limiting for polymerase activity. Stu2 likely operates under a similar regime in cells, because the concentration of tubulin in yeast has been estimated to be on the order of 0.5 μM (Wethekam and Moore, 2023), which is comparable to the concentrations used in our in vitro experiments. Thus, Stu2 functions like a tubulin shuttling antenna that enhances capture of unpolymerized tubulin and accelerates its incorporation into the microtubule end.

Nothing in the biochemical model specifies how the tubulin-binding TOG domains need to be linked beyond that they be linked flexibly. Thus, the model should not be limited to homodimeric polymerases such as Stu2: it should apply equally well to monomeric polymerases that contain different numbers of TOG domains such as Zyg9 (3 TOGs) or XMAP215 and chTOG (5 TOGs). For any given polymerase, the overall activity will in general be determined by several factors. The number of tubulin-binding TOG domains and how many polymerases accumulate on the microtubule end together dictate the total number of TOG domains available to shuttle/deliver tubulin. The affinity and kinetics of TOG:tubulin interactions will determine the proportional contributions to polymerase activity from different TOGs, with lower affinity or slower binding TOGs contributing proportionally less than higher affinity, faster binding TOGs (e.g. (Widlund et al., 2011)). In addition to the polymerase-intrinsic factors above, whether a given polymerase is affinity or kinetically limited will also depend on the concentration of tubulin, which influences the rates of tubulin:TOG and tubulin:microtubule binding.

In summary, we showed that the same enzyme-like biochemical model can explain the activity of otherwise unrelated microtubule and actin polymerases: each polymerase resides at the polymer end and functions like a shuttling antenna, using multiple flexibly-linked actin or tubulin binding domains to capture and accelerate delivery of unpolymerized subunits to the polymer end. For the Stu2/XMAP215 microtubule polymerases, this is to our knowledge the first time that the polymerase activity has been quantitatively explained in terms of the number of TOG domains and their tubulin-binding rate constants. That two otherwise unrelated cytoskeletal polymerases share the same mechanism provides an interesting example of convergent evolution.

Materials and Methods

Plasmid construction

Plasmids expressing wild-type αβ yeast tubulin as well as polymerization blocked tubulin, LR1 (β: T175R,V179R) were previously described (Ayaz et al., 2012; Johnson et al., 2011). For sortase mediated peptide ligation, a sortase recognition sequence (LPETGG) was added to polymerization blocked tubulin at its C-terminus. The Stu2 variants - TOG1-TOG2, TOG2-TOG2, and TOG1*-TOG2 - were expressed using the pHAT vector, which includes an N-terminal HIS tag, a C-terminal eGFP sequence and Strep-tag, as previously described (Geyer et al., 2015; Geyer et al., 2018). Stu2-TOG2 (318–560) expressed from pET15b with a N-terminal HIS tag (Ayaz et al., 2012). All constructs were confirmed by DNA sequencing.

Protein expression and purification

Wild-type and polymerization-blocked αβ-tubulin were purified from inducibly overexpressing strains of Saccharomyces cerevisiae using Ni-affinity and anion exchange chromatography as described previously (Ayaz et al., 2012; Johnson et al., 2011). Purified tubulin samples were stored in 10 mM K-PIPES pH 6.9, 2 mM MgSO4, 1 mM EGTA with 50 µM GTP. Stu2 variants (TOG1-TOG2, TOG2-TOG2, TOG1*-TOG2) were overexpressed in E. coli using Arctic Express Cells using 0.5 mM IPTG for 24 hrs at 12 °C. Cell pellets were resuspended in lysis buffer (50 mM sodium phosphate dibasic and monobasic mix, 300 mM NaCl, 40 mM imidazole, 5% glycerol), PMSF was added to 1 mM final concentration and cells were lysed using a microfluidizer. Crude lysate was clarified by centrifugation before loading onto a His60 Superflow Column (Clontech). Column-bound proteins were eluted in Ni-elution buffer (lysis buffer + 300 mM imidazole). Pooled elution fractions were loaded onto a Strep-Tactin Superflow column (IBA, Germany) and eluted in Strep-elution buffer (RB100 + 10 mM desthiobiotin). Eluted samples were buffer exchanged into RB100 with 2 mL, 7K MWCO Zeba spin desalting columns (Thermo Scientific) for storage. Stu2-TOG2 (318-560) was overexpressed in E. coli and purified using a combination of Ni affinity and cation exchange chromatography as described previously (Geyer et al., 2015). Eluted samples were buffer exchanged into RB100 (25 mM Tris pH 7.5, 100 mM NaCl, 1 mM MgCl2, 1 mM EGTA) or dynamics assay buffer (90 mM PIPES, 10 mM NaCl, 1 mM MgCl2, 1 mM EGTA, 50 µM GTP), depending on the assays and flash frozen.

In vitro microtubule dynamics assay

Microtubule dynamics were imaged by either differential interference contrast microscopy (DIC) or total internal reflection fluorescence (TIRF) microscopy as previously described (Geyer et al., 2018). Briefly, assay chambers were assembled from slides, coverslips, and double sticky tape. The chambers were incubated with 5 mg/ml neutravidin (Sigma) and blocked using 1% F-127 Pluronic acid for 10 mins each. The chambers were then rinsed with BRB80 (80 mM PIPES pH 6.9, 1 mM MgCl2, 1 mM EGTA) followed by 10 min incubation with GMPCPP seeds made from brain tubulin (5% biotinylated, PurSolutions). Chambers were rinsed again with BRB80 and samples containing tubulin, Stu2 in 1X PEM (90 mM K-PIPES, 1 mM EGTA, 1 mM MgSO4) + 0.1 mg/ml bovine serum albumin (BSA) + 1 mM GTP were flowed into the same and immediately sealed with VALAP. Microtubule dynamics were measured manually by creating kymographs with ImageJ (Edelstein et al., 2010) and analyzed as previously described (Geyer et al., 2018)

End intensity measurements

To measure if there is a steady amount of polymerase at the growing MT end, the MT were imaged by TIRF. Flow chambers were prepared as stated in the previous section with addition of antifade agents (glucose, glucose oxidase, and catalase). Stu2 concentration was held constant at 100 nM while WT tubulin was used at 0.6μM and 1.4μM.

All imaging conditions such as exposure time, imaging angle were kept as identical as possible between two ranges of tubulin. Kymographs were created in ImageJ. For a given kymograph, intensities were measured at three different points of a growing MT (Fig. 3A) by drawing a uniform length line across each of those points and using the plot profile feature in ImageJ. The maximum and minimum gray values (intensity) for each plot were manually subtracted and analyzed using GraphPad Prism 10.2.

Bio-layer interferometry

We used sortase-mediated protein splicing to selectively label polymerization-blocked tubulin at the C-terminus of β-tubulin. GGGC peptide (Genscript) was incubated with a 20-fold molar excess of EZ-Link™ Maleimide-PEG2-Biotin (MPB) at room temperature. The reaction was quenched with 100 mM DTT after 2 hours. This biotinylated peptide provides the substrate for sortase-mediated labeling of tubulin. Labeling of polymerase-blocked tubulin was carried at room temperature for 30 minutes, with 50-fold molar excess of GGGC-MPB and 1.6-fold molar excess of sortase to polymerization-blocked tubulin, in sortase buffer (20 mM Tris pH 7.5, 125 mM NaCl, 1 mM TCEP, 0.01 mM CaCl₂ and 50 nM GTP). The reaction product was purified again using ion exchange chromatography to get rid of free peptides. The labeled blocked tubulin was confirmed using mass spectrophotometry.

BLI binding measurements were performed using Streptavidin (SA) biosensors on an OctetR8 instrument (Sartorius, Germany). The assay buffer consisted of 90 mM PIPES, 10 mM NaCl, 1 mM MgCl₂, 1 mM EGTA, 1 mM GTP, 0.1% BSA and 0.005% triton X-100. (BSA and triton X-100 were added to reduce non-specific binding to the biosensors). Biotinylated polymerization-blocked tubulin was immobilized on SA biosensors, which were then washed and dipped into wells containing TOG2 at concentrations ranging from 2.3 to 200 nM. The response level during immobilization was kept below 1 nm to minimize avidity-like artifacts. The association step lasted 300 seconds and was followed by a dissociation step, where the biosensor was moved into a well containing assay buffer, lasting 300 seconds. Data were double reference-subtracted using the signal from an unloaded pin dipped into TOG2 and a pin loading with LR1-457 dipped into assay buffer without TOG2. Association and dissociation rates were determining using a 1:1 binding mode analysis with the Octet BLI Discovery 13.0 (Orthwein et al., 2021).

Mathematical model for Stu2 mediated microtubule elongation

In this study, we adapted the Ena/VASP model for Stu2, as previously described (Breitsprecher et al., 2011). Briefly, the saturation dependence curve for tubulin shown in Figure 2D is reminiscent of the Michaelis-Menten model (V=V_max[S]/K_M+[S]) for enzyme kinetics with an assumption that each growing MT tip is associated with a few TOG domains, N. These domains are responsible for capturing tubulin with a rate constant k_on and transferring it onto the MT growing end with rate k_t. The tubulin can also be released back into the solution with an off rate k_off. Briefly, when we fit the MT elongation data with a Michaelis-Menten model, K_M corresponds to K_D+k_t/k_on and V_max corresponds to k_tN. By solving for the apparent K_M using K_D and k_on, we estimated k_t, which was then used to determine the value of ‘N’ for different Stu2 variants. The data were analyzed using GraphPad Prism 10.2.

Acknowledgements

We thank E. Bonventre, R. Chen, and L. McCormick for critical feedback on the manuscript. This work was supported by grants from the NIH: R01GM098543 and R35GM156385 to LMR, and R00GM141261 and R01GM155152 to DLK.