Mechanism of Microtubule Stabilization by Doublecortin

Molecular Cell, Vol. 14, 833–839, June 18, 2004, Copyright 2004 by Cell Press

Mechanism of Microtubule Stabilization by Doublecortin Carolyn A. Moores,1,4 Myle`ne Perderiset,2 Fiona Francis,3 Jamel Chelly,3 Anne Houdusse,2 and Ronald A. Milligan1,* 1 Department of Cell Biology, CB227 The Scripps Research Institute 10550 North Torrey Pines Road La Jolla, California 92037 2 Motilite´ Structurale Institut Curie CNRS UMR 144 26 rue d’Ulm 75248 Paris Cedex 05 France 3 Laboratoire de Ge´ne´tique et Physiopathologie des Retards Mentaux, GDPM Institut Cochin 24 rue du Faubourg Saint Jacques Paris, 75014 France

Summary Neurons undertake an amazing journey from the center of the developing mammalian brain to the outer layers of the cerebral cortex. Doublecortin, a component of the microtubule cytoskeleton, is essential in postmitotic neurons and was identified because its mutation disrupts human brain development. Doublecortin stabilizes microtubules and stimulates their polymerization but has no homology with other MAPs. We used electron microscopy to characterize microtubule binding by doublecortin and visualize its binding site. Doublecortin binds selectively to 13 protofilament microtubules, its in vivo substrate, and also causes preferential assembly of 13 protofilament microtubules. This specificity was explained when we found that doublecortin binds between the protofilaments from which microtubules are built, a previously uncharacterized binding site that is ideal for microtubule stabilization. These data reveal the structural basis for doublecortin’s binding selectivity and provide insight into its role in maintaining microtubule architecture in maturing neurons. Introduction An extraordinary level of orchestrated cellular movement occurs during the development of the human cerebral cortex (Gupta et al., 2002). Billions of immature neurons migrate out of the inner regions of the developing brain and form the six precisely organized layers of the mature cerebral cortex (Olson and Walsh, 2002). As with other cell types, cytoskeletal elements are cen*Correspondence: [email protected] 4 Current address: Department of Crystallography, Birkbeck College, University of London, Malet Street, London WC1E 7HX, United Kingdom.

Short Article

tral to neuronal motility and differentiation (Feng and Walsh, 2001). However, perhaps because of the complexity of the pathfinding that neurons must perform, because of the relatively long distances traveled, or because many precise cell-cell contacts must ultimately be formed, they also require neuron-specific molecules. The existence of these molecules is demonstrated most vividly when their function is knocked out by genetic mutation, either by direct manipulation in experimental animals or in human disease (Couillard-Despres et al., 2001). One such disease is lissencephaly, where the six well-organized cellular layers of the cerebral cortex are replaced by four poorly ordered layers (Gleeson, 2000) and results in epilepsy and mental retardation in lissencephaly patients. One of the genes whose mutation can lead to lissencephaly is the X-linked doublecortin gene (des Portes et al., 1998; Gleeson et al. 1998). Its 40 kDa protein product, doublecortin (DCX), is expressed in migrating and differentiating neurons and is associated with the microtubule (MT) cytoskeleton (Francis et al., 1999; Gleeson et al., 1999), consistent with its essential role in neuronal maturation. DCX can be divided into two functional parts (Figure 1A): the N-terminal 30 kDa part is the MT binding portion of the protein and is composed of two homologous, 11 kDa DC domains. The importance of MT binding for DCX’s function in the neuron is shown by the clustering of lissencephalycausing mutations within these DC repeats (Sapir et al., 2000; Taylor et al., 2000). However, little is known about this binding interaction, and despite the homology of the DC repeats, it is not clear whether these domains have equivalent functionalities with respect to their interaction with MTs (Kim et al., 2003). The Ser/Pro-rich C-terminal portion of the protein is not required for MT binding, and an interaction site for clathrin-associated proteins and for several protein kinases has been localized to this domain (Friocourt et al., 2001; Gdalyuhu, et al., 2004; Tanaka et al., 2004). DCX binds MTs in vivo and in vitro and stimulates MT polymerization, thus sharing many properties with the well-characterized neuronal MAPs, including tau and MAP2 (Cassimeris and Spittle, 2001). However, DCX does not share any sequence homology with these proteins and, in contrast to the random coil structure of classical neuronal MAPs (Butner and Kirschner, 1991), the DC domains of DCX adopt globular structures with a ubiquitin-like fold (Kim et al., 2003). To understand how DCX binds MTs and stabilizes them, we used cryoelectron microscopy to examine the interactions of DCX with MTs and to locate the DCX binding site. Results Doublecortin Binds Specific Microtubule Architectures In this study, we used recombinant full-length doublecortin (DCX) and a truncated version consisting of the N-terminal 30 kDa containing the two DC repeats

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but lacking the S/P-rich C-terminal domain (t-DCX, Figure 1A; see Experimental Procedures). The truncated version is sufficient for MT binding both in vivo and in vitro (Horesh et al., 1999; Sapir et al., 2000; Taylor et al., 2000; Kim et al., 2003). We bound these DCX constructs to in vitro polymerized, taxol-stabilized MTs and examined the resultant complexes by cryo-electron microscopy. Although it was difficult to see DCX decoration directly in the images, regular binding by both constructs was evidenced by the presence of an 80 A˚ signal in optical diffraction patterns of individual MTs. Undecorated MTs do not have an 80 A˚ signal, and its appearance demonstrates that the stoichiometry of binding is one DCX per tubulin heterodimer. In preliminary experiments, we observed that DCX binding to MTs was variable; within the same image, the protein bound to some MTs (80 A˚ diffraction evident) but not to others (no 80 A˚ diffraction). On closer inspection, we found that binding correlated with the number of protofilaments from which the MT was constructed (Figure 1B). The truncated construct, t-DCX, preferentially bound to MTs with low protofilament (pf) number. Under the conditions of the study, ⵑ50% of MTs with 12 pf were decorated with t-DCX, whereas only ⵑ5% of 15 pf MTs showed any evidence of binding, and no evidence of binding to 16 pf MTs was seen. Full-length doublecortin, DCX, was even more substrate specific, showing a distinct preference for 13 pf MTs and a total lack of binding to either 15 or 16 pf MTs (Figure 1B). As most cellular MTs, and in particular those within the neuron, are built from 13 pfs (Tilney et al., 1973; Moritz and Agard, 2001), the data suggest that DCX has evolved specifically to recognize this architecture. Furthermore, although the N-terminal DC domains of t-DCX target smaller diameter (lower pf number) MTs, additional specificity is conferred by the C-terminal S/P-rich domain found in the full-length protein. Tubulin—Doublecortin Copolymerization Results in 13 pf Microtubules As doublecortin has been shown to enhance the polymerization of pure tubulin (Horesh et al., 1999; Sapir et al., 2000; Taylor et al., 2000), we next asked if it had any effect on the outcome of MT polymerization. In vitro polymerization of pure tubulin resulted in a range of MT architectures (Wade et al., 1990; Ray et al., 1993): ⵑ70%

Figure 1. Doublecortin Selectively Binds 13 Protofilament Microtubules and Nucleates Microtubules (A) Domain organization of doublecortin. N-DC and C-DC each comprise 85 residues. The C-terminal Ser/Pro-rich domain is made up of 100 residues, the majority of which is missing in the t-DCX construct used to determine the doublecortin binding site. (B) t-DCX (gray bars) and DCX (white bars) show binding selectivity

for specific MT architectures. Plotted values are percent total MTs of a given pf number that showed an 80 A˚⫺1 signal in the optical diffractometer. Total number of MTs examined was 361 and 224 for t-DCX and DCX decoration, respectively. (C) MTs copolymerized with doublecortin have predominantly 13 pfs (white bars). Polymerization of pure tubulin results in mostly 14 pf MTs (black bars) (total polymer length, 354 ␮m and 353 ␮m, respectively). For cryo-electron microscopy, 25 ␮M tubulin was copolymerized with 34 ␮M full-length DCX at 37⬚C for 6 hr prior to freezing. (D) Typical negative stain image of the result of tubulin polymerization in the absence (top panel) and presence (bottom panel) of DCX. Scale bar, 1 ␮m. (E) Average length of MTs polymerized in the absence (⫺DCX) and presence (⫹DCX) of full-length doublecortin, compared to MTs polymerized with GMPCPP (measured from negative stain EM images).

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14 pf, ⵑ20% 13 pf, and some 12 pf, 15 pf, and 16 pf microtubules (Figure 1C). The effect of copolymerization with DCX was dramatic, abolishing the formation of 15 and 16 pf MTs and greatly reducing the number of 14 and 12 pf MTs. Approximately 70% of the MTs grown in the presence of DCX had 13 pfs (Figure 1C). Thus, not only does DCX selectively bind to 13 pf MTs, it also leads to growth of predominantly 13 pf MTs. Doublecortin Is a Microtubule Nucleator Another striking observation following MT copolymerization with DCX was its effect on MT length. When the MT products of DCX copolymerization were compared with MTs polymerized in the absence of DCX under identical conditions, the ⫹DCX MT were shorter (Figure 1D). With pure tubulin, polymerization is a two step process in which tubulin heterodimers must first form a nucleation structure and then add to and elongate this MT nucleus (Job et al., 2003). At a given tubulin concentration, if nucleation is favored then lots of short MTs will be seen. On the other hand, if growth is favored, fewer longer MTs will form, and thus, our results support a role for DCX in MT nucleation. In addition, the average length of MTs polymerized with DCX was comparable to that seen with MTs polymerized in the presence of the nonhydrolyzable GTP analog GMPCPP (Figure 1E), which is a potent MT nucleator (Hyman et al., 1992). Although more work is required to characterize the kinetics of DCX copolymerization, our observations support the idea that DCX stimulates MT nucleation and that this nucleation results in 13 pf MTs. Doublecortin Binds between Microtubule Protofilaments We next used cryo-electron microscopy and helical image analysis to locate the site of DCX binding on MTs. As 13 pf MTs are not helical and DCX binds very poorly to 14 pf MTs (see Figure 1B), we analyzed helical 14 pf MTs that were decorated with the truncated construct t-DCX. We calculated 3D maps from images of undecorated and t-DCX-decorated MTs and located t-DCX by difference mapping (see Experimental Procedures). This analysis revealed strong difference peaks of high statistical significance lying 80 A˚ apart, in one of the two structurally distinct fenestrations between adjacent pfs

Figure 2. Doublecortin Binds in the Valley between Microtubule Protofilaments (A) Front view of the 3D difference map (yellow) representing t-DCX, displayed with a 3D map of the undecorated MT (blue). The difference map was obtained by subtracting the 3D map of the undecorated MT from a 3D map of t-DCX-decorated MT. Doublecortin binds between tubulin pfs and lies over one of the two distinct fenestrations in the MT wall. (B) View from the plus end showing how doublecortin fits snugly into the valley between adjacent pfs. Orange and green areas ap-

proximately delineate the kinesin and MAP2/tau binding sites respectively. The asterisk shows the location of the taxol binding site. (C) Close-up view of (A), with the atomic model of two tubulin heterodimers (Lo¨we et al., 2001) docked into the MT map. The yellow density attributable to doublecortin lies at the junction of four tubulin monomers (labeled ␣1, ␤1, ␣2, and ␤2). We emphasize that our data do not allow us to tell whether doublecortin binds at the location shown—between two heterodimers or at the similar but distinct site 40 A˚ away—at the junction between four heterodimers (asterisk). (D) View of (C) from the MT plus end shows that doublecortin makes relatively sparse contacts with individual monomers, suggesting that its binding site is unique to polymerized tubulin. (E) The doublecortin difference map can only accommodate a single DC domain (left). The right panel shows N-DC in an equivalent orientation and illustrates that surface residues mutated in lissencephaly (R78, D86, R89, R102, shown in space-filling representation) map to one side of the DC domain and are likely involved in MT binding. (Scale bars, 20 A˚).

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in the MT wall (Figure 2A). The protein appears to be wedged into the valley between adjacent pfs and does not overlap the binding sites for kinesin or MAP2/tau (Figure 2B) (Al-Bassam et al., 2002; Moores et al., 2003). Docking the atomic structure of tubulin dimers (Lo¨we et al., 2001) into the MT portion of the map shows clearly that t-DCX binds at the junction of four tubulin monomers (Figures 2C and 2D). We cannot distinguish between ␣- and ␤-tubulin in our 3D maps, so the t-DCX density is shown over the fenestration between two adjacent tubulin heterodimers for purposes of illustration only. As is most apparent in the view from the MT plus end (Figure 2D), t-DCX does not seem to contact any major elements of tubulin secondary structure. Instead, it binds in an environment unique to polymerized tubulin that is created by loops from four tubulin subunits (H9␤8 loop in ␣1, H10-␤9 loop in ␤1, H11-H12 loop in ␣2, H4-␤5 loop in ␤2). Although ␣- and ␤-tubulin are structurally very similar, the sequences are only 45% identical in the loop regions where t-DCX binds. The sequence differences represent a likely signature by which DCX can distinguish between the similar but not identical fenestrations in the valleys of the outer surface of the MT wall and target to the correct one. To extend the insights gained from the difference mapping, we made use of the recently solved highresolution structure of the N-terminal DC domain of DCX (Kim et al., 2003). The DC core of this structure and the difference density are of comparable size, suggesting that we have visualized only one of the two DC domains of t-DCX (Figure 2E). We believe our difference density represents the N-terminal DC domain since this domain selectively binds MTs (Kim et al., 2003), whereas the second, more C-terminal, DC domain has also been implicated in binding free tubulin and other cellular partners (Caspi et al., 2000; Tsukada et al., 2003). There is some additional low-level density in our 3D map located over the difference peak (data not shown), suggesting that the second DC domain may be present but disordered or poorly localized at higher MT radius. The 40 residue linker sequence between the two DC domains may contribute to poor localization of the second domain. That only one DC domain is clearly revealed by our analysis is however consistent with biochemical characterization of the different contributions N-DC and C-DC make to the overall properties of DCX, even though both domains are required for full functionality (Taylor et al., 2000). Our data may represent a snapshot of what is probably a range of conformational and functional states for C-DC; for example, its interaction with tubulin dimers may be important as MT nucleation is initiated and as polymerization proceeds, but a welldefined interaction involving this domain may not be required in the fully polymerized MTs analyzed here. Lissencephaly-causing mutations in DCX have been classified according to whether they disrupt the DC fold, thereby abolishing function, or are located on the protein surface, where they presumably disrupt the interaction with MTs or other proteins (Kim et al., 2003). The t-DCX difference map does not provide sufficient information to identify a unique orientation for N-DC within the electron density. However, the clustering of disease-causing mutations (R78, D86, R89, R102; Figure 2E) on one face of N-DCX strongly suggests that this face is functionally

important and likely to be involved in binding the tubulin loops described above (Figure 2E). This docked orientation for N-DC provides a testable model for future experiments. Discussion Modulation of Microtubule Dynamics MT dynamics is an essential attribute of the cytoskeleton in migrating neurons and growth cones (GordonWeeks, 2004) with MT dynamic instability governed by the guanine nucleotide bound to ␤-tubulin in the tubulin heterodimer (Desai and Mitchison, 1997). GTP-tubulin heterodimers adopt a conformation that favors their incorporation into straight pfs and enables lateral contacts (inter-pf) to form the MT wall. GDP-tubulin dimers have a curved conformation and, if not held in place by the MT lattice, will favor depolymerization through outward curling of the pfs, concomitant with loss of lateral contacts (Ravelli et al., 2004). Based on these ideas, it is clear that molecules that stabilize straight pfs and/or strengthen interactions between adjacent pfs will reduce depolymerization and increase MT stability. Among the proteins that bind along MTs and affect dynamics, the MAP2/tau family is perhaps the best studied (Cassimeris and Spittle, 2001). These molecules are unstructured in solution but bind in an ordered conformation along the crest of the MT protofilament (Figure 2B). Although the precise mechanism of MT stabilization by MAP2/tau is not known, it appears to be a consequence of the extended molecule spanning and interacting with the interdimer and intradimer interfaces along the pf crests and maintaining the straight protofilament conformation (Al-Bassam et al., 2002). In contrast, Taxol威—another potent MT stabilizer— can be thought of as an agent that strengthens lateral contacts between pfs (although allosteric effects on longitudinal contacts cannot be excluded). Taxol binds on the inside wall of the MT close to the site of interaction between ␤-tubulins in adjacent pfs (asterisk in Figure 2B; Nogales et al., 1999). It is thought to stabilize structural elements that are predicted to be conformationally variable depending on the phase of dynamic instability (Amos and Lo¨we, 1999). Selective Binding of Doublecortin to 13 pf Microtubules The data we have presented here now expand this repertoire of stabilization strategies to include one in which the mechanism is particularly well matched to the type of MT occurring in vivo. Our structural study reveals DCX density—most likely the N-DC domain—wedged into the valley between pfs. Given that the hinge-like contacts between pfs are located toward the inside of the MT wall (see Figure 2D), purely geometrical considerations dictate that the width of the valley is dependent on pf number, with wider valleys in low pf MTs and narrower valleys in high pf MTs. The t-DCX construct has an increasing propensity to bind MTs as their pf number decreases (Figure 1B), suggesting that simple accessibility of the binding site plays a major role in controlling the interaction—the wider the valley, the easier it is for the protein to access its binding site. However,

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when the C-terminal S/P domain is present (Figure 1B), the simple accessibility mechanism must be refined to include a degree of specificity, with the S/P domain responsible for selecting (directly or indirectly) valley dimension in a narrow range. In contrast to the results of binding MAP2/tau proteins or taxol—which are independent of pf number—DCX and, presumably, its relatives (Burgess and Reiner, 2000; Lin et al., 2000) appear to be extremely well tuned to interact with the 13 pf MTs found in vivo. We only see what is presumably the N-DC domain clearly in our maps, and the additional weak density at higher radius (data not shown) may be attributable to a poorly localized C-DC domain. Since both DC domains are required for full functionality of DCX (Horesh et al., 1999; Taylor et al., 2000), poor localization of the C-DC domain, together with the observation that it can also bind tubulin heterodimers (Taylor et al., 2000; Kim et al., 2003), raises the possibility that C-DC may participate in initial nucleation steps and subsequent recruitment and positioning of incoming tubulin at the ends of growing pfs. Along the length of stabilized MTs, however, it will be available to interact with other cellular ligands (Kizhatil et al., 2002; Tsukada et al., 2003). In addition, even though DCX’s C-terminal, S/P-rich domain is required for proper targeting to 13 pf MTs, its accessibility to other cellular binding partners is likely to be critical in the integration of cell signaling and cytoskeletal dynamics during neuronal migration and differentiation (Friocourt et al., 2001; Gdalyuhu et al., 2004; Tanaka et al., 2004). Microtubule Stabilization Both lateral (between pfs) and longitudinal (along each pf) interactions between tubulins act together to stabilize the cylindrical MT structure. Although DCX binding between pfs is an excellent way to crosslink pfs together and thus increase MT stability, it is noteworthy that DCX’s position within the valley suggests the possibility of additional mechanistic sophistication. By binding over a fenestration, which may be the weakest point in the MT wall, DCX interacts with four tubulin monomers. In doing so, it not only has the potential to strengthen the lateral connections between pfs but is also in an ideal position, by virtue of its interaction with two adjacent tubulin monomers along each pf, to augment longitudinal interactions. Thus, as well as “stapling” pfs together around the MT wall, DCX may act at the tubulin junctions along pfs to maintain the straight conformation, in contrast to MAP2/tau, which seems to act only along pfs. Strikingly, a similar stapling mechanism was recently reported for the actin-stabilizing protein SipA (Lilic et al., 2003). Microtubule Nucleation The mechanism of MT nucleation from pure tubulin in vitro is not well understood (Job et al., 2003) but results in MTs with a range of pf number (Figure 1C; Wade et al., 1990; Ray et al., 1993). Our copolymerization experiments suggest that DCX may stimulate nucleation and that it affects the outcome of assembly, resulting in predominantly 13 pf MTs. Given the location and geometrical constraints of the DCX binding site on the MT,

we envision two possible mechanisms that could explain this outcome. First, DCX may accelerate the formation of initial nucleation seeds and subsequently constrain wall closure so that 13 pf geometry is favored. In this model, 13 pf MTs are preferentially nucleated and stabilized. Alternatively, accelerated nucleation of MTs with a variety of pf number may occur, but since DCX binds preferentially to 13 pf MTs, these are stabilized and removed from the dynamic pool of assembling MTs. In this model, the outcome is a result of selective stabilization of a particular architecture in a dynamic population of assembling MTs. Cellular Implications of Doublecortin Action Directed MT polymerization is an essential aspect of cell motility, including neuronal migration and pathfinding (Feng and Walsh, 2001). Typically, MT nucleation occurs at the centrosome, where ␥-TuRCs act as templates for growth of 13 pf MTs (Evans et al., 1985; Moritz and Agard, 2001). Neurons face a unique set of challenges in organizing their MT array because of the distance between the centrosome and their most distal processes. They solve this problem by releasing MTs from the centrosome and funnel them toward their extremities (Dent et al., 1999; Gordon-Weeks, 2004; and references therein). Local MT dynamics are central to neuronal growth cone movement (Buck and Zheng, 2002), and short dynamic MTs are found distally in the processes of migrating neurons in regions where DCX is concentrated (Friocourt et al., 2003; Schaar et al., 2004). Thus, the abilities of DCX to bind and stabilize MTs within these processes and to be regulated by extracellular cues are ideal traits for a protein involved in brain development. There is some evidence for centrosomal-independent nucleation in nonneuronal cells (Vorobjev et al., 1997; Yvon and Wadsworth, 1997), but it is not known whether such local nucleation events occur in neuronal growth cones. If they do, DCX may function to ensure that 13 protofilament MTs are preferentially nucleated. MT stabilization by DCX could also facilitate the formation and maintenance of the MT cage around the nucleus of migrating neurons (Tanaka et al., 2004). Another possibility is that DCX may exert a corrective influence during polymerization, ensuring that 13 pf architecture is maintained. Rapidly elongating microtubules can sometimes change pf number (Chretien et al., 1992). If this happens in cells, the selective stabilization model described above would act to correct the mistake and ensure that only the desired 13 pf architecture is stabilized. Nevertheless, the properties of DCX that we have described here make it clear that it can reinforce and stabilize the 13 pf geometry of prepolymerized MTs in these cells by an unusually well-designed mechanism distinct from that of classical neuronal MAPs. DCX’s selectivity for 13 pf MTs serves to underscore the significance of this polymer architecture in the majority of cell types (Tilney et al., 1973). The reason that this architecture is desirable is unclear. The pfs of these MTs lie parallel to the MT axis and do not have the supertwist of MTs with other pf numbers (Ray et al., 1993). This design may have evolved to provide the shortest route from one end of a MT to the other for both growth and MT-based transport. In the context of

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the entire cell, these factors may make a significant contribution to cytoskeleton efficiency and organization. In addition, MT bundling by crosslinking proteins may be accomplished more effectively in MTs with untwisted pfs. Finally, uninterrupted, processive, motor-based transport could in principle occur on the outer surface of 13 pf MT bundles; transport on bundles of supertwisted (non-13 pf) MTs would be interrupted as the pf tracks rotate away from the surface of the bundle. Irrespective of the underlying significance of 13 pf architecture in MTs in vivo, it seems clear that DCX has evolved to ensure that it is maintained. The accumulation of DCX at the leading edge of motile and immature neurons and its kinase-dependent regulation here (Gdalyuhu et al., 2004; Schaar et al., 2004) is a functionally significant location for its role in migration and differentiation and is distinct from that of other MAPs (Francis et al., 1999; Friocourt et al., 2003). The properties of DCX revealed by our binding and structural studies provide insight into the failure of neuronal MAPs to compensate when DCX is absent and further our understanding of its essential role in neuronal motility and brain development. Experimental Procedures Doublecortin Expression The cDNA for the two DCX constructs were ligated to the pFastBac HTa baculovirus donor plasmid (Bac-to-Bac Baculovirus Expression System, Life Technologies). The full-length construct DCX consisted of 6xHIS-rtTEV-NcoI-FLAG-BglII-doublecortin 1:366-STOP-EcoRISstI-XbaI. t-DCX consisted of 6xHIS-rtTEV-NcoI-FLAG-BglIIdoublecortin 1:282-STOP-SphI-KpnI-HindIII. The transfer vector containing each doublecortin construct was transfected into Spodoptera frugigerpa (Sf9) using the Bac-to-Bac Baculovirus Expression System. Sf9 cells in suspension were infected for 48 hr with viral stock at various multiplicities of infection. Doublecortin Purification Cells were harvested in PBS and lysed in a buffer containing 50 mM HEPES, 20 mM imidazole, 300 mM NaCl, 0.5% Triton, 5 mM ␤-mercaptoethanol, 0.5 mM PMSF, 1 ␮g/ml leupeptin (pH 7.5). Cell debris was spun down (100,000 ⫻ g, 30 min), and doublecortin was purified from the supernatant by Ni-NTA affinity chromatography. The His6-tag was removed by tobacco etch virus (TEV) protease. Doublecortin was further purified by gel filtration chromatography in 25 mM PIPES, 100 mM NaCl, 5 mM Tris (2-carboxyethyl) phosphine hydrochloride (TCEP) (pH 6.8), and concentrated to 6 mg/ml. MT Polymerization For electron microscopy and structure determination, bovine brain tubulin (Cytoskeleton Inc.) at a final concentration of 5 mg/ml was polymerized in 80 mM PIPES (pH 6.8), 1.5 mM MgCl2, 8% DMSO, and 2.5mM GTP for 30 min at 35⬚C, after which 1 mM paclitaxel (Cal-Biochem) dissolved in DMSO was added. To characterize the effect of DCX on MT polymer length, MTs were polymerized at 37⬚C using the buffer conditions described above with the following modifications: (1) 15 ␮M full-length DCX was added at the beginning of polymerization of 10 ␮M tubulin and (2) MTs were polymerized in the presence of GMPCPP instead of GTP. The different MT preparations were left at room temperature for 24 hr. MTs were adsorbed to carbon-coated grids and were negatively stained with 1% uranyl acetate. The method used for estimation of MT length is given in Supplemental Data at http://www.molecule.org/cgi/content/full/14/ 6/833/DC1. Preparation of Frozen Grids for Cryo-Electron Microscopy Taxol-stabilized MTs at 1.25 mg/ml were applied to glow-discharged 400 mesh Quantifoil grids with 2 ␮m holes in a carbon support film

(Signal Probe Co.). The grids were washed with BrB80, and then, t-DCX dialyzed against BrB80 ⫹ 1 mM TCEP was applied to the grid. The grids were blotted and frozen by plunging them rapidly into liquid ethane slush (Dubochet et al., 1988) and were stored under liquid nitrogen. Cryo-electron microscopy and helical image processing was performed as described in Moores et al. (2003), the details of which are given in Supplemental Data available on Molecular Cell’s website. Difference Mapping and Atomic Model Docking A difference map was calculated between the ⫾ t-DCX 3D maps, and the statistical significance of differences was calculated using a Student’s t test (Milligan and Flicker, 1987). ␣␤-tubulin dimer atomic coordinates (Lo¨we et al. 2001; PDB# 1JFF) and human N-DC atomic coordinates (Kim et al., 2003; PDB# 1MJD) were manually docked into the t-DCX-decorated MT map and were rendered using AVS (Advanced Visual Systems, Inc.). The position of the N-DC lissencephaly mutations was rendered using PyMOL (Delano Scientific, LLC). Acknowledgments We thank T. Mitchison (Harvard Medical School) for the gift of GMPCPP. This work was supported by grants from the National Institutes of Health (GM52468, GM61939, and RR17573), the European Commission (QLG3-CT-2000-00158), INSERM, CNRS, the French Ministe`re de la Recherche (ACI-1A066G), the Fe´de´ration pour la Recherche sur le Cerveau, the Fondation pour la Recherche Me´dicale (20000293/3-INE), and the Association pour la Recherche sur le Cancer (ARC 5505) (to A.H.). Received: February 27, 2004 Revised: April 26, 2004 Accepted: April 28, 2004 Published: June 17, 2004 References Al-Bassam, J., Ozer, R.S., Safer, D., Halpain, S., and Milligan, R.A. (2002). MAP2 and tau bind longitudinally along the outer ridges of microtubule protofilaments. J. Cell Biol. 157, 1187–1196. Amos, L.A., and Lo¨we, J. (1999). How Taxol stabilizes microtubule structure. Chem. Biol. 6, R65–R69. Buck, K.B., and Zheng, J.Q. (2002). Growth cone turning induced by direct local modification of microtubule dynamics. J. Neurosci. 22, 9358–9367. Burgess, H.A., and Reiner, O. (2000). Doublecortin-like kinase is associated with microtubules in neuronal growth cones. Mol. Cell. Neurosci. 16, 529–541. Butner, K.A., and Kirschner, M.W. (1991). Tau protein binds to microtubules through a flexible array of distributed weak sites. J. Cell Biol. 115, 717–730. Caspi, M., Atlas, R., Kantor, A., Sapir, T., and Reiner, O. (2000). Interaction between Lis1 and doublecortin, two lissencephaly gene products. Hum. Mol. Genet. 15, 2205–2213. Cassimeris, L., and Spittle, C. (2001). Regulation of microtubuleassociated proteins. Int. Rev. Cytol. 210, 163–226. Chretien, D., Metoz, F., Verde, F., Karsenti, E., and Wade, R.H. (1992). Lattice defects in microtubules: protofilament numbers vary within individual microtubules. J. Cell Biol. 117, 1031–1040. Couillard-Despres, S., Winkler, J., Uyanik, G., and Aigner, L. (2001). Molecular mechanisms of neuronal migration disorders, quo vadis? Curr. Mol. Med. 1, 677–688. Dent, E.W., Callaway, J.L., Szebenyi, G., Baas, P.W., and Kalil, K. (1999). Reorganization and movement of microtubules in axonal growth cones and developing interstitial branches. J. Neurosci. 19, 8894–8908. Desai, A., and Mitchison, T.J. (1997). Microtubule polymerization dynamics. Annu. Rev. Cell Dev. Biol. 13, 83–117. des Portes, V., Pinard, J.M., Billuart, P., Vinet, M.C., Koulakoff, A., Carrie´, A., Gelot, A., Dupuis, E., Motte, J., Berwald-Netter, Y., et al.

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