The kinesin-13 MCAK has an unconventional ATPase cycle adapted for microtubule depolymerization
2011; Springer Nature; Volume: 30; Issue: 19 Linguagem: Inglês
10.1038/emboj.2011.290
ISSN1460-2075
AutoresClaire T. Friel, Jonathon Howard,
Tópico(s)Photosynthetic Processes and Mechanisms
ResumoArticle26 August 2011Open Access The kinesin-13 MCAK has an unconventional ATPase cycle adapted for microtubule depolymerization Claire T Friel Corresponding Author Claire T Friel Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, GermanyPresent address: School of Biomedical Sciences, University of Nottingham, Medical School, Queen's Medical Centre, Nottingham NG7 2UH, UK. Tel.: +44 115 82 30138; E-mail: [email protected]Search for more papers by this author Jonathon Howard Corresponding Author Jonathon Howard Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany Search for more papers by this author Claire T Friel Corresponding Author Claire T Friel Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, GermanyPresent address: School of Biomedical Sciences, University of Nottingham, Medical School, Queen's Medical Centre, Nottingham NG7 2UH, UK. Tel.: +44 115 82 30138; E-mail: [email protected]Search for more papers by this author Jonathon Howard Corresponding Author Jonathon Howard Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany Search for more papers by this author Author Information Claire T Friel 1 and Jonathon Howard 1 1Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany *Corresponding authors. Max Planck Institute of Molecular Cell Biology and Genetics, Pfotenhauerstr. 108, 01307 Dresden, Germany. Tel.: +49 351 210 2500; Fax: +49 351 210 2020Max Planck Institute of Molecular Cell Biology and Genetics, Pfotenhauerstr. 108, 01307 Dresden, Germany. Tel.: +49 351 210 2500; Fax: +49 351 210 2020; E-mail: [email protected] The EMBO Journal (2011)30:3928-3939https://doi.org/10.1038/emboj.2011.290 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Unlike other kinesins, members of the kinesin-13 subfamily do not move directionally along microtubules but, instead, depolymerize them. To understand how kinesins with structurally similar motor domains can have such dissimilar functions, we elucidated the ATP turnover cycle of the kinesin-13, MCAK. In contrast to translocating kinesins, ATP cleavage, rather than product release, is the rate-limiting step for ATP turnover by MCAK; unpolymerized tubulin and microtubules accelerate this step. Further, microtubule ends fully activate the ATPase by accelerating the exchange of ADP for ATP. This tuning of the cycle adapts MCAK for its depolymerization activity: lattice-stimulated ATP cleavage drives MCAK into a weakly bound nucleotide state that reaches microtubule ends by diffusion, and end-specific acceleration of nucleotide exchange drives MCAK into a strongly bound state that promotes depolymerization. This altered cycle accounts well for the different mechanical behaviour of this kinesin, which depolymerizes microtubules from their ends, compared to translocating kinesins that walk along microtubules. Thus, the kinesin motor domain is a nucleotide-dependent engine that can be differentially tuned for transport or depolymerization functions. Introduction The kinesins are a superfamily of proteins defined by a common, highly conserved motor domain (Marx et al, 2009). For most kinesins, the motor domain functions as a molecular machine that converts chemical energy derived from the turnover of ATP into mechanical work used for directed motion along microtubules (Vale and Milligan, 2000; Howard, 2001; Schliwa, 2003). Directed motion is used to translocate organelles (kinesin-1, kinesin-2 and kinesin-3) (Hirokawa et al, 2009) or chromosomes (kinesin-4 and kinesin-10) (Mazumdar and Misteli, 2005) along microtubules, or to slide antiparallel arrays of microtubules (kinesin-5 and kinesin-14) (Peterman and Scholey, 2009). In contrast to the transport and sliding kinesins, members of the kinesin-13 family do not move in a directed manner on microtubules. Instead, they bind weakly to the microtubule lattice and undergo random, diffusive motion (Helenius et al, 2006; Cooper et al, 2009). The motor function of kinesin-13s occurs at microtubule ends where, in a reaction coupled to ATP turnover, they catalytically depolymerize the stable GTP-containing caps of growing microtubules (Desai et al, 1999; Hunter et al, 2003; Wagenbach et al, 2008). Loss of the GTP cap, which can be mimicked using the slowly hydrolyzed analogue GMPCPP, converts the growing microtubule into a shrinking one (catastrophe). This depolymerase activity accounts well for the cellular phenotypes caused by disruption of kinesin-13. For example, immunodepletion of kinesin-13 from frog egg extracts leads to enlarged mitotic spindles (Walczak et al, 1996) and injection of antibodies to kinesin-13 blocks microtubule dynamics in the Drosophila early embryo (Rogers et al, 2004). Conversely, overexpression of kinesin-13 in tissue culture cells leads to an increase in the frequency of catastrophes (Kline-Smith and Walczak, 2002) and a reduction in the number of microtubules (Maney et al, 1998), again consistent with kinesin-13 proteins being depolymerases in vivo. Thus, kinesin-13s have very different activities to most other kinesins. This is remarkable given the high sequence and structural conservation of the kinesin motor domain (Ogawa et al, 2004; Shipley et al, 2004). That motor domains of similar structure confer such different functions suggests that the internal workings of kinesin-13 differ from those of other kinesins. All kinesins studied to date, including kinesin-13s, use nucleotide turnover to alternate between states of high and low microtubule affinity: without nucleotide and in the presence of the non-hydrolyzable ATP analog, AMPPNP, kinesins bind strongly to microtubules, whereas in the presence of ADP they bind weakly. For the translocating kinesins, this change in affinity is coupled to directed movement of the motor domain along the microtubule. In the kinesin-1 family, the release of ADP is the rate-limiting step in the absence of microtubules and is accelerated ∼1000-fold by microtubule binding (Hackney, 1988). Following microtubule binding and release of ADP, the binding of ATP induces a conformational change in the switch regions resulting in the neck linker docking towards the plus end of the microtubule (Sindelar and Downing, 2010). For dimeric kinesin-1, this facilitates binding of the second head to the next site on the microtubule (Hackney, 1994a; Schief et al, 2004), allowing processive hand-over-hand movement, tightly coupled to ATP hydrolysis (Kaseda et al, 2002; Asbury et al, 2003; Yildiz et al, 2004). The conventional cycle of ATP turnover outlined in the previous paragraph makes no sense for kinesin-13s, which do not translocate along the lattice. Instead, the motor activity of kinesin-13s occurs at the microtubule end (Desai et al, 1999; Hunter et al, 2003; Moore and Wordeman, 2004), where the end-stimulated ATPase of MCAK allows catalytic microtubule depolymerization. Previous work has shown that ATP cleavage is not required for the tubulin removal step (Desai et al, 1999; Moores et al, 2002; Wagenbach et al, 2008), suggesting that it is necessary for another aspect of catalytic activity, such as recycling of MCAK from tubulin heterodimers (Desai et al, 1999; Hertzer and Walczak, 2008). To understand how the ATP turnover cycle of a kinesin may be tuned to produce a microtubule depolymerase, we have elucidated the ATPase cycle of the kinesin-13 MCAK (mitotic centromere associated kinesin) in solution and in the presence of unpolymerized tubulin and microtubules. The cycle differs in fundamental ways from other kinesins and these differences tailor MCAK to its function as a microtubule depolymerase. Results The low basal ATPase of MCAK is accelerated by tubulin and microtubules The key question is which step (or steps) in the ATP hydrolysis cycle of MCAK is rate limiting, and how this rate-limiting step (or steps) is altered by tubulin and microtubules. To answer this question, we first measured the rate constant for the complete ATP turnover cycle: the rate of product formation per motor domain (ATPase rate). The basal ATPase rate of MCAK (the ATPase rate without tubulin or microtubules) was calculated by measuring the rate of formation of the reaction product, inorganic phosphate (Pi; Figure 1A) and dividing by the concentration of MCAK motor domains (each MCAK molecule has two ATP-hydrolyzing motor domains). The basal ATPase rate of MCAK is very low, 0.00095±0.00005 s−1 (mean±s.d., n=3). A similar low rate was determined by measuring the production of the other reaction product, ADP (Supplementary Figure S1A). The ATPase rate was increased by unpolymerized tubulin to 0.123±0.005 s−1 (mean±s.e.) and by microtubules to 4.97±0.53 s−1 (mean±s.e.) at saturating tubulin and microtubules (Table I, Supplementary Figure S1B and C). Thus, unpolymerized tubulin accelerates the ATPase ∼100-fold and microtubules ∼5000-fold. Figure 1.MCAK utilizes mantATP equally well as unlabelled ATP. (A) Production of inorganic phosphate during the reaction of MCAK with ATP (open circles) or mantATP (red squares). Linear fit to the data ATP (black) and mantATP (red). The basal ATPase rate determined by this method was 0.00095±0.00005 s−1 for ATP and 0.00090±0.00003 s−1 for mantATP (mean±s.d., n=3). (B) Dependence of the ATPase rate for MCAK upon the concentration of mantATP. Fit to the Michaelis–Menton equation (black line): Vmax=0.00085±0.00003 s−1, KM=0.40±0.08 μM (mean±s.e.). (C) Kymographs of fluorescently labelled GMPCPP-stabilised microtubules depolymerized by MCAK in the presence of 1 mM ATP or mantATP. Addition of MCAK is indicated by the red arrowhead. The depolymerization rate was 0.58±0.16 μm min−1 (mean±s.d., n=16) and 0.55±0.14 μm min−1 (mean±s.d., n=14) for ATP and mantATP, respectively. (D) Distribution of depolymerization rates for individual microtubules upon the addition of 100 nM MCAK plus 1 mM ATP or mantATP. The box represents the central 50% of the distribution, the central line the median and the whiskers the full range of the distribution. Download figure Download PowerPoint Table 1. Compiled rate constants for the ATP turnover cycle of MCAK (mean±s.e.) Dissection of the ATP turnover cycle using fluorescent nucleotides The ATP turnover cycle comprises four individual chemical steps (): ATP binding to the empty site (denoted by ϕ) (1), ATP cleavage (2), phosphate dissociation (3) and ADP dissociation (4). To determine the rate-limiting step, we used nucleotide labelled with the fluorescent moiety mant (methylanthraniloyl) (Supplementary Figure S2A). The small mant group positioned on the ribose has been shown to have no significant effect on the binding and hydrolysis of nucleotides by several ATP-hydrolyzing enzymes (Sadhu and Taylor, 1992; Gilbert et al, 1995; Lisal and Tuma, 2005). Likewise, MCAK can use mantATP as a substrate: the basal mantATPase rate for MCAK was 0.00090±0.00003 s−1 (mean±s.d., n=3), not significantly different to the basal unlabelled ATPase rate (Figure 1A). The mantATPase rate depended upon the concentration of mantATP (Figure 1B) giving a KM=0.40±0.08 μM (mean±s.e.). The Vmax of 0.00085±0.00003 s−1 (mean±s.e.) is in agreement with the basal mantATPase rate measured by monitoring the production of inorganic phosphate. Importantly, in the presence of mantATP, MCAK depolymerized microtubules at the same rate as in the presence of unlabelled ATP (Figure 1C and D). To determine the association and dissociation rate constants for ATP (, k1 and k−1) in the absence of tubulin or microtubules, the interaction of nucleotide-free MCAK (the bound ADP is removed by incubating with excess EDTA (Sadhu and Taylor, 1992)) with mantATP was assayed by rapidly mixing, using stopped-flow, and measuring the time course of the change in fluorescence. Upon binding to MCAK, the fluorescence of the mant group increased approximately two-fold (Figure 2A). When the concentration of mantATP was in excess of the MCAK concentration, the observed increase in fluorescence was well described by a single exponential, plus a line of constant negative slope to account for photobleaching of the mant group. The rate constant associated with the exponential increased linearly with the concentration of mantATP (Figure 2B). The gradient of the linear fit to these data provided the association rate constant (k1=0.164±0.007 μM−1 s−1, mean±s.e.) and the intercept provided the dissociation rate constant (k−1=0.041±0.001 s−1, mean±s.e.) for mantATP (Table I). Figure 2.Binding of mantATP and dissociation of mantADP. (A) Fluorescence increase upon mixing mantATP (3.1 μM) with nucleotide-free (ϕ) MCAK (0.6 μM) using stopped-flow. The fluorescence signal (black) is fit to a single exponential function, plus a line of constant negative slope to account for photobleaching of the mant group (red dashed). Inset: Contents of the syringes prior to mixing. (B) Dependence of the rate constant for the interaction of mantATP with nucleotide-free MCAK upon the concentration of mantATP (open circles). The gradient of the linear fit (red) gives the rate constant for mantATP binding, 0.164±0.007 μM−1 s−1 (mean±s.e.), and the intercept the mantATP dissociation rate constant, 0.041±0.001 s−1 (mean±s.e.). (C) Fluorescence decrease upon the dissociation of mantADP from MCAK. The fluorescence signal (black) is fit to a single exponential function, plus a line of constant negative slope to account for photobleaching of the mant group (red dashed). Inset: contents of the syringes prior to mixing, MCAK preloaded with mantADP is mixed with an excess of unlabelled ATP, which prevents rebinding of mantADP. Download figure Download PowerPoint An increase in fluorescence was also observed for mantADP binding to nucleotide-free MCAK (Supplementary Figure S2B). The fit to these data provided the rate constants for association (k−4=0.85±0.11 μM−1 s−1, mean±s.e.) and dissociation (k4=0.16±0.03 s−1, mean±s.e.) of mantADP (Supplementary Figure S2C). ADP dissociation is not the rate-limiting step in MCAK's basal hydrolysis cycle The result in the previous paragraph suggests that ADP dissociation from MCAK is about 150 times faster than the overall cycle rate (0.16 s−1 compared to 0.00095 s−1). To confirm this, we directly measured the rate constant for ADP dissociation (, k4) by preloading MCAK with mantADP and rapidly mixing with an excess of unlabelled ATP. The fluorescence of mantADP decreased as it dissociated from MCAK and was replaced by the unlabelled nucleotide (Figure 2C). The time course of this decrease in fluorescence was well described by a single exponential, when corrected for photobleaching of the mant group. The rate constant was 0.126±0.015 s−1 (mean±s.d., n=15) (Table I) consistent with the earlier indirect measurement. As this rate constant is so much larger than the basal ATPase rate, ADP dissociation cannot be the rate-limiting step for MCAK. This distinguishes MCAK from translocating kinesins. A highly fluorescent intermediate during the reaction of MCAK with mantATP ATP binding is fast relative to the ATPase rate: Figure 2B gives a rate constant of ∼4 s−1 at 25 μM ATP. Thus, neither ATP binding nor ADP release is rate limiting. Therefore, either ATP cleavage or the release of inorganic phosphate, Pi (, Steps 2 and 3), must be rate limiting. To investigate these steps, we used low turnover conditions (Supplementary Information), in which the concentration of nucleotide is lower than that of the protein. This allowed us to observe intermediates transiently populated during the ATPase cycle. Again, the use of mant-labelled nucleotides allowed us to resolve individual steps in the cycle. Addition of 1 μM ADP·MCAK (each MCAK motor domain has a bound ADP unless specifically treated to remove this nucleotide) to a solution containing 0.8 μM mantATP resulted in an unusual biphasic change in fluorescence (Figure 3A, black curve). Following the addition of ADP·MCAK (indicated by the red arrowhead, Figure 3A), the mant fluorescence first increased and then decreased. The rising phase corresponds to displacement of ADP by mantATP (, Steps 4 and 1), resulting in the previously observed increase in fluorescence of mantATP upon binding to MCAK. We attribute the slower, falling phase to the conversion of mantATP to mantADP in the nucleotide-binding pocket (, Steps 2 and 3). This interpretation is based on the following arguments. First, the fluorescence intensity at the end point of the reaction is similar to the fluorescence intensity obtained when ADP·MCAK is added to mantADP (Figure 3A, blue curve). Second, the addition of excess unlabelled ATP at the end point of the reaction (Supplementary Figure S2D) led to a fluorescence decrease with the same rate constant (0.114±0.012 s−1) as that previously measured (0.126±0.015 s−1) for the dissociation of mantADP from MCAK. An alternative explanation is that the biphasic kinetics result from differing affinity of mantATP for the individual motor domains of the MCAK dimer. However, biphasic kinetics are also observed for a monomeric MCAK construct (Figure 3B). We conclude that mantATP·MCAK has a higher fluorescence signal than mantADP·MCAK; this has been observed for other motor proteins (Woodward et al, 1991; Ma and Taylor, 1995; Kuhlman and Bagshaw, 1998). Figure 3.ATP cleavage is rate limiting in the ATP turnover cycle of MCAK. (A) Fluorescence change upon the reaction of 1 μM ADP·MCAK with 0.8 μM mantATP (black). A highly fluorescent intermediate (red asterisk) is observed during the reaction. Single exponential fit to the phase of decreasing fluorescence (red dashed), k=0.00068±0.00026 s−1 (mean±s.d., n=8). Reaction of 1 μM ADP·MCAK with 0.8 μM mantADP (blue). Reactions were initiated by manual addition of ADP·MCAK (red arrowhead) to a solution of mantATP or mantADP. The fluorescence signal is normalized to the mant-nucleotide signal prior to addition of MCAK. Inset: Reaction of 1 μM ADP kinesin-1 (Drosophila melanogaster KHC 1-557) with 0.8 μM mantATP. Black arrowhead indicates addition of the kinesin. (B) Fluorescence change upon mixing monomeric ADP·MCAK (NM-MCAK, A181-V585) with a substoichiometric concentration of mantATP. Inset: syringe contents prior to mixing. (C) mantATP as a fraction of total nucleotide at the indicated time after the initiation of the reaction of 1.0 μM ADP·MCAK with 0.5 μM mantATP (open circles). Fit of these data to a single exponential function (red), k=0.00045±0.00009 s−1 (mean±s.e.). Inset: example of HPLC traces showing the change in the proportions of mantATP (T) and mantADP (D) as the reaction progresses. The fraction of ATP remaining was determined by dividing the area under the peak corresponding to mantATP by the sum of the area under the mantATP and the mantADP peaks. (D) Time course of the formation of mantADP upon the reaction of 1 μM nucleotide-free MCAK with 10-fold excess mantATP. The ATPase rate was 0.0018±0.0007 (mean±s.d., n=3). The dashed line indicates the magnitude of the burst of product that would be observed if product release (, Step 3 or 4) rather than ATP cleavage (, Step 2) was rate limiting. Download figure Download PowerPoint The biphasic change in fluorescence during the reaction of ADP·MCAK with mantATP confirms that ADP release is not the rate-limiting step in this experiment. If it were, then the reaction would be monophasic with a rate constant equal to the rate constant of ADP dissociation. Such a monophasic curve is seen for kinesin-1 (Figure 3A, inset); the rate constant of 0.02 s−1 is similar to that previously reported (Hackney, 1988). The rate constant determined for the phase of decreasing fluorescence was 0.00068±0.00026 s−1 (mean±s.d., n=8) (Table I), less than but of the same order of magnitude as the basal mantATPase rate (0.00090±0.00003 s−1). The rate constant, associated with the fluorescence decrease, is consistent with the rate-limiting step in the basal ATP turnover cycle of MCAK. The rate constant of the fluorescence decrease is expected to be lower than the basal ATPase rate because not all of the mantATP binds to MCAK in the initial binding event (Supplementary Information). Given the affinity for mantATP (Figure 2B) and the presence of ADP equimolar with MCAK motor domains, we estimate that the rate constant of the fluorescence decrease should be ∼44% of the ATPase rate (Supplementary Information). We conclude that either the cleavage of ATP or the dissociation of phosphate (, Step 2 or 3) represents the rate-limiting step in the basal ATP turnover cycle of MCAK. ATP cleavage is rate limiting for the ATP hydrolysis cycle of MCAK in solution The rate-limiting step is either ATP cleavage or Pi dissociation. To distinguish between these two possibilities, we identified the highly fluorescent intermediate (Figure 3A, red asterisk). We took samples of the reaction over time, quenched the reaction to release the protein-bound nucleotides and separated them on a C18 column (Figure 3C, inset). As the reaction must be carried out under low turnover conditions ([ATP]<[MCAK]), low concentrations of ATP must be used. Therefore, we used mantATP as the substrate, enabling fluorescence detection of the nucleotides giving greater sensitivity. The fraction of mantATP remaining was determined over the course of the reaction (Figure 3C). The fraction of mantATP decreased exponentially with a rate constant of 0.00045±0.00009 s−1 (mean±s.e.) (Table I). This is not statistically significantly different (by a t-test) from the rate constant of the phase of decreasing fluorescence in Figure 3A (0.00068) and corresponds to ∼50% of the basal mantATPase rate (0.00090 s−1). The agreement of these rate constants indicates that the highly fluorescent intermediate seen in Figure 3A is mantATP. Thus, ATP cleavage, not phosphate release, is the rate-limiting step in the conversion of mantATP to mantADP (, Step 2). To confirm the conclusion that ATP cleavage rather than Pi dissociation is rate limiting, the kinetics of ATP turnover were monitored by quenching the reaction of mantATP with MCAK and measuring the total amount of product (both free and bound to the enzyme) over time. Nucleotide-free MCAK was used at a concentration more than double the KM to ensure substrate binding was not limiting. We observed a linear increase in product (Figure 3D) with a mantATPase rate of 0.0018±0.0007 (mean±s.d., n=3). This rate is similar to, though somewhat higher than, the ATPase rates measured in earlier assays (mantATPase=0.00090 s−1, by Pi production (Figure 1A) and 0.00085 s−1, by mantADP production (Figure 1B); unlabelled ATPase=0.00095 s−1, by Pi production (Figure 1A) and 0.0017 s−1, by ADP production (Supplementary Figure S1A)). The important point is that there is no burst of product, confirming that ATP cleavage is the rate limiting step in MCAK's ATP turnover cycle (Foster and Gilbert, 2000). Unpolymerized tubulin and microtubules accelerate ATP cleavage Both unpolymerized and polymerized tubulin (microtubules) increase the ATPase rate. Therefore, they must both accelerate ATP cleavage. To determine the identity of the rate-limiting step in the presence of unpolymerized tubulin and microtubules, we monitored the reaction of mantATP with ADP·MCAK, under low turnover conditions, by rapid mixing using stopped-flow (Figure 4A). Without tubulin or microtubules (Figure 4B, upper trace) a biphasic fluorescence transient was observed, as for the same reaction initiated by manual mixing (Figure 3A). The addition of 0.5 μM unpolymerized tubulin accelerated the second phase (Figure 4B, middle trace, black arrow). Addition of 10 μM unpolymerized tubulin, which saturates the ATPase (Supplementary Figure S1B) and ensures that tubulin binding is not rate limiting, accelerated the second phase to such an extent that it was no longer observable (Figure 4B, lower trace). Thus, in the presence of saturating tubulin concentrations, ATP cleavage is no longer the rate-limiting step. Figure 4.Unpolymerized tubulin or microtubules change the rate-limiting step in MCAK's ATP turnover cycle. (A–C) The reaction of ADP·MCAK with a substoichiometric concentration of mantATP, alone or in the presence of unpolymerized tubulin or microtubules, initiated by rapid mixing using a stopped-flow fluorimeter. (A) Syringe contents prior to mixing: ADP·MCAK is mixed 1:1 v/v with mantATP plus unpolymerized tubulin or microtubules as stated. (B) Fluorescence change upon the reaction of 0.5 μM ADP·MCAK with 0.4 μM mantATP in the absence or presence of 0.5 μM or 10 μM unpolymerized tubulin (post-mixing concentrations). (C) Fluorescence change upon the reaction of 0.5 μM ADP·MCAK with 0.4 μM mantATP in the presence of 5 μM tubulin polymer (post-mixing concentrations). In (B) and (C), the fluorescence is normalized to the respective initial signals. (D) Sequential mixing assay in which ADP·MCAK is premixed with a substoichiometric concentration of mantATP, incubated for 30s (i.e. until the reaction has progressed to the point marked by the asterisk in B, upper trace) and then mixed with microtubules. (E) Fluorescence change upon mixing 0.5 μM ADP·MCAK pre-incubated for 30 s with 0.4 μM mantATP with microtubules. In panels B, C and E the data are shown in black and the fit to the data in red. Download figure Download PowerPoint In the presence of 10 μM unpolymerized tubulin, the rate constant of the increase in fluorescence (Figure 4B, lower trace) was 0.250±0.022 s−1 (mean±s.d., n=4) (Table I). We attribute this rate constant to the dissociation of ADP from MCAK: it is similar to the rate constant for mantADP dissociation in the absence (0.126±0.015 s−1) and presence (0.128±0.017 s−1; mean±s.d., n=4) of tubulin (Table I, Supplementary Figure S3A and B). Taken together, these data indicate that unpolymerized tubulin increases the ATPase rate for MCAK by increasing the rate constant for the cleavage of ATP, such that ADP dissociation becomes rate limiting for the ATP turnover cycle. Microtubules accelerate both ATP cleavage and ADP dissociation The addition of 5 μM polymerized tubulin to the reaction of ADP·MCAK with mantATP dramatically changed the kinetics of the reaction. The phase of decreasing fluorescence, seen for the reaction without tubulin or microtubules, was no longer visible (Figure 4C, compare with Figure 4B, upper trace). Instead, the fluorescence increased monotonically. The absence of the phase of decreasing fluorescence, previously shown to report on the conversion of mantATP to mantADP (, Steps 2 and 3), indicates that these steps occur faster than displacement of ADP by mantATP (, Steps 4 and 1). Thus, neither cleavage of mantATP nor dissociation of phosphate is rate limiting in the presence of microtubules. To confirm the identity of the process responsible for the observed fluorescence increase and thus the rate-limiting step for the cycle, these data were compared to the reaction of mantADP·MCAK with unlabelled ATP, which reports solely on ADP dissociation (, k4). The fluorescence decrease observed (Supplementary Figure S3C) occurred with the same rate constants as the fluorescence increase observed in Figure 4C. Therefore, we conclude that the rate-limiting step for ATP turnover in the presence of microtubules is ADP dissociation. The microtubule-stimulated dissociation of ADP (Figure 4C) had two phases: a larger, faster phase (amplitude 72–79%) with a rate constant of 5.17±0.41 s−1 (mean±s.d., n=4) (Table I); and a smaller, slower phase (amplitude 21–28%) with a rate constant of 0.19±0.06 s−1 (mean±s.d., n=4). We attribute the faster phase to the dissociation of ADP from MCAK, accelerated ∼35-fold by microtubules, as this rate constant is similar to the ATPase rate in the presence of microtubules, 4.97±0.53 s−1 (Table I). These data indicate that ADP release is rate limiting for microtubule-stimulated ATP turnover, and is accelerated relative to the same process in solution or in the presence of unpolymerized tubulin. The smaller, slow phase (Figure 4C) has a rate constant similar to ADP dissociation from MCAK in the presence of unpolymerized tubulin (Table I, Supplementary Figure S3B), which may be due to unpolymerized tubulin present in the sample of microtubules binding to ADP·MCAK and sterically blocking microtubule-stimulated ADP release. Alternatively, it may be due to MCAK binding to some very long microtubules in the reaction mix and failing to reach the end, where ADP release is maximally accelerated (see below). Irrespective of its origin, this phase is small and too slow to represent the rate-limiting step of the microtubule-stimulated ATPase. The conclusion that the conversion of mantATP to mantADP is accelerated in the presence of microtubules is based on the absence of the phase of decreasing fluorescence (compare Figure 4C with Figure 4B, upper trace). To confirm that the conversion of mantATP to mantADP is accelerated by microtubules, the order of mixing in the assay in Figure 4C was altered (Figure 4D) to allow visualization of the associated change in fluorescence. ADP·MCAK was pre-incubated for 30 s with mantATP, allowing mantATP to bind to the MCAK motor domain. The preformed mantATP·MCAK was then mixed with microtubules (Figure 4E). A rapid decrease in fluorescence was observed, with a rate constant of 10.04±0.54 s−1 (mean±s.d., n=3). Thus, the conversion of mantATP to mantADP in the presence of microtubules occurs ∼14 000-fold faster t
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