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1 Department of Molecular Cell Biology, Institute of Molecular Embryology and Genetics, Kumamoto University, 2-2-1 Honjo, Kumamoto 860-0811, Japan
2 International Graduate School of Arts and Sciences, Yokohama City University, 1-7-29 Suehiro-cho, Tsurumi-ku, Yokohama 230-0045, Japan
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
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Introduction |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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Sequence analyses revealed that spastin is composed of the AAA ATPase domain in the C-terminal region and the MIT (microtubule interacting and trafficking) domain and MTBD (microtubule-binding domain) in the N-terminal region, suggesting an active role in cytoskeleton interactions (Takasu et al. 2005; White et al. 2007). MTBD has been identified in human spastin to be necessary for microtubule association and severing (White et al. 2007). On the other hand, the MIT domain of Vps4p plays an important role in the interaction with ESCRT-III (endosomal sorting complexes required for transport III) proteins and is definitely required for its function, multivesicular body formation and viral budding (Obita et al. 2007; Stuchell-Brereton et al. 2007). The MIT domain is also found in a number of proteins involved in endosomal function (Scott et al. 2005a). The MIT domain of Drosophila spastin has also been shown to facilitate ATPase activity, and microtubule-binding and -severing activities (Roll-Mecak & Vale 2008).
AAA proteins are considered to remodel substrates: unfolding and disassembly of proteins and protein complexes (Vale 2000; Ogura & Wilkinson 2001; Lupas & Martin 2002; White & Lauring 2007). AAA proteins generally oligomerize into a hexamer and there is the pore in the hexameric ring (Ogura & Wilkinson 2001; Yakushiji et al. 2006; White et al. 2007; Pantakani et al. 2008; Roll-Mecak & Vale 2008). It has been suggested that the central pore is a channel through which substrates are translocated (Voges et al. 1999; Kim et al. 2000; Ogura & Wilkinson 2001; Yamada-Inagawa et al. 2003). Structural studies of AAA proteins revealed that the highly conserved aromatic residue in the AAA domain lies in the central pore region of hexamer (Yamada-Inagawa et al. 2003; Schlieker et al. 2004; Gerega et al. 2005; Scott et al. 2005b). This conserved pore residue is important for the function. In the AAA protease FtsH of Escherichia coli, when the conserved pore residue was mutated, mutant FtsH proteins abolished proteolytic activity (Yamada-Inagawa et al. 2003).
On the other hand, Vps4p/SKD1, which is classified into the meiotic subfamily containing spastin, katanin and fidgetin, dissociates ESCRT complexes from membranes during multivesicular body formation and viral budding (Babst et al. 1998; Obita et al. 2007; Stuchell-Brereton et al. 2007). Three other AAA proteins, katanin, spastin and fidgetin, disassemble microtubules during chromosome segregation, and axon outgrowth and branch formation (McNally & Vale 1993; Errico et al. 2004; Zhang et al. 2007). It has been reported that katanin assembles to oligomers on microtubules in an ATP-dependent manner, and suggested that ATP hydrolysis may change the conformation of the katanin ring, leading to mechanical strain that destabilized tubulin–tubulin contacts (Hartman & Vale 1999). However, it has also been demonstrated that the pore mutant of Vps4p blocked viral budding (Scott et al. 2005b). Moreover, it has been suggested that the hexameric form of spastin recognizes the C-terminal region of 
/β-tubulin, and pull or translocate these peptides through central pore of the hexamer (White et al. 2007; Roll-Mecak & Vale 2008). Thus, mechanism of unfolding and disassembly is in dispute.
Our previous analyses on the mutant of Caenorhabditis elegans spastin homologue SPAS-1 suggested that C. elegans SPAS-1 is involved in microtubule dynamics (Matsushita-Ishiodori et al. 2007). When expressed in the cultured cells, microtubule disassembly was observed as same as observed with spastin from human, mouse and fly (Evans et al. 2005; Roll-Mecak & Vale 2005; Matsushita-Ishiodori et al. 2007; Solowska et al. 2008). In this study, we found that SPAS-1 formed a stable hexamer in a concentration-dependent manner. Given that MTBD of SPAS-1 is essential for binding to tubulin, we propose that MTBD of SPAS-1 plays a critical role in enrichment of SPAS-1 to microtubules, where SPAS-1 is concentrated and able to form a stable hexamer. Furthermore, the mutational analyses revealed that the conserved solvent exposed, basic amino acid residues in the pore region of spastin as well as the well-conserved aromatic residue are important for the recognition and severing of microtubules.
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Results |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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The entire open reading frame for SPAS-1 was cloned into the His-tag expression vector pET-15b as described in Experimental procedures. From this construct, SPAS-1 protein (451 amino acid residues) was produced as an N-terminally His6-tagged recombinant protein. SPAS-1 was recovered mostly in a soluble fraction after ultracentrifugation (data not shown). After purified with Ni2+-NTA agarose, SPAS-1 was greater than 90% pure as judged from SDS–PAGE analysis (Fig. 1a).
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-helical structure (Fig. 1b). Next, to test for thermal transition of the secondary structure, we monitored conformational transitions of SPAS-1 as the solution temperature increased (from 25 to 95 °C). CD spectra showed the decrease in ellipticity at 208 and 222 nm, suggesting that the content of -helices decreased dramatically by heating (Fig. 1b). The melting temperature Tm was estimated as 47 °C from the denaturation curve (Fig. 1c). To demonstrate whether SPAS-1 possesses an ATPase activity, purified SPAS-1 (2.2 µg) was incubated with ATP at 37 °C for 30 min. ATPase activity of 70 nmol/mg/min was obtained (Fig. 1d). It is well known that mutations in the Walker A or B motif of AAA proteins abolish ATPase activity. To eliminate the possibility that the observed activity is due to the contamination, we prepared these mutant proteins (Walker A mutant SPAS-1K224R and Walker B mutant SPAS-1E278Q). As shown in Fig. 1d, ATPase activities were nearly completely abolished by these mutations, indicating that the purified SPAS-1 protein indeed has an ATPase activity. Effects of pH, salt, Mg2+and DTT on ATPase activity were subsequently examined. The optimum condition for ATPase activity of SPAS-1 was thus determined to be as follows: the reaction is carried out in 10 mM Tris–HCl (pH 8.8), 50 mM KCl, 5 mM Mg(CH3COO)2, 5 mM DTT and 3 mM ATP in a 25-µL reaction mixture at 37 °C. With varying ATP concentrations from 0.3 to 3 mM, Lineweaver–Burk plot was made (data not shown). Km and Vmax for the ATPase activity of SPAS-1 were estimated to be 1.53 mM and 115 nmol/µg/min, respectively. These values are comparative with a Km of 0.33 mM and Vmax of 520 nmol/µg/min for the recombinant p97/VCP (Song et al. 2003), with a Km of 0.44 mM and Vmax of 225 nmol/µg/min for the recombinant C. elegans fidgetin (Yakushiji et al. 2004), with a Km of 0.27 mM and Vmax of 660 nmol/µg/min for the recombinant human spastin (Evans et al. 2005), and with a Km of 1.497 mM and Vmax of 287 nmol/µg/min for the recombinant human spastin (Pantakani et al. 2008). As discussed previously (Song et al. 2003; Yakushiji et al. 2004), it is consistent with the notion that AAA family members all possess weak ATPase activities.
In general, substrates stimulate the ATPase activity of AAA proteins (Hartman et al. 1998; Makyio et al. 2002; Yamada-Inagawa et al. 2003; Azmi et al. 2006). As spastin homologues target microtubules (Evans et al. 2005; Roll-Mecak & Vale 2005; Wood et al. 2006; Matsushita-Ishiodori et al. 2007; Zhang et al. 2007), we examined the effects of microtubules on the SPAS-1 ATPase activity. It was found that SPAS-1 displays a complex stimulation pattern by microtubules (Fig. 1e). SPAS-1’s ATPase activity increased with increasing concentrations of microtubules up to 0.25 µM and then declined at higher microtubule concentrations. In the presence of 0.25 µM microtubules, ATPase activity was seven times higher than that obtained in the absence of microtubules (Fig. 1e). These results may imply that microtubules stimulate the ATPase activity of SPAS-1 by facilitating SPAS-1–SPAS-1 interactions, and that high concentrations of microtubules reduce the ATPase activity by preventing SPAS-1–SPAS-1 associations through the sequestration of SPAS-1 monomers, as demonstrated in the case of katanin (Hartman & Vale 1999). In contrast, stimulation of the ATPase activity of FtsH by its substrates casein and 
32 displays typical hyperbolic curves that reach saturation (Yamada-Inagawa et al. 2003).
Oligomer formation of SPAS-1
AAA proteins are generally considered to form a hexamer, whose formation is critical for their function (Shotland et al. 1997). It has been demonstrated that spastin assembles into a hexamer in an ATP-dependent (White et al. 2007; Roll-Mecak & Vale 2008) or -independent (Pantakani et al. 2008) manner. We tested whether purified C. elegans SPAS-1 forms a hexamer or not. It contains a His6-tag sequence at N-terminus as described in Experimental procedures and therefore the molecular size of monomer is c. 52 kDa. When highly concentrated SPAS-1 (>5 mg/mL) was analyzed by size exclusion column chromatography, SPAS-1 hexamers were detected (Fig. 1f). It should be mentioned that the buffer used for chromatography did not contain ATP. Hence, to decipher the ATP requirement for hexamer formation, ATP-binding deficient mutant SPAS-1K224R and ATP-hydrolysis deficient mutant SPAS-1E278Q were analyzed. As shown in Fig. 1f, hexamers were observed as same as those observed with wild-type SPAS-1, implying that the hexamer formation of SPAS-1 takes place in an ATP-independent manner. It is interesting to note that the formation of oligomer with molecular mass of c. 820 kDa was more prominent in SPAS-1E278Q (see Discussion). By contrast, when low concentrations of SPAS-1 (<0.55 mg/mL) were analyzed, stable hexamers were not formed, rather it shows a broad elution profile over monomer to hexamer, even in the presence of ATP (Fig. 1g). These results together indicate that the oligomerization of SPAS-1 is concentration dependent, but does not require ATP.
SPAS-1 directly interacts with tubulin
It has been reported that spastin is involved in microtubule severing, and the microtubule-binding domain was identified as MTBD (Evans et al. 2005; Roll-Mecak & Vale 2005; White et al. 2007). We also demonstrated that C. elegans SPAS-1 showed microtubule-severing activity when expressed in cultured cells (Matsushita-Ishiodori et al. 2007). We therefore examined if SPAS-1 binds to tubulin. When /β-tubulin dimer was incubated with SPAS-1, tubulin was pulled down with SPAS-1 (Fig. 2a), suggesting that C. elegans SPAS-1 also binds to tubulin. Next, to determine the binding domain of SPAS-1 to tubulin, we prepared a series of nested deletions of His6-SPAS-1, and performed pull-down assay. While SPAS-11–210 and SPAS-11–152 bound to tubulin as similar as full-length SPAS-1 did, SPAS-11–103 showed no ability to interact with tubulin (Fig. 2a), suggesting that residues 103–152 are necessary for interaction with tubulin and that the MIT domain alone does not interact with tubulin.
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/β-tubulin dimer more rigorously using surface plasmon resonance analysis. As clearly shown in Fig. 2b, SPAS-11–152 bound the immobilized /β-tubulin and response units increased proportionally with increasing concentrations of SPAS-11–152 with a dissociation constant of a KD of 0.24 µM, indicating that the interaction between SPAS-1 and tubulin is direct and specific to SPAS-11–152. In addition, we prepared SPAS-1115–172 containing the region corresponding to MTBD of human spastin, which has been identified as a MTBD (White et al. 2007), and performed pull-down assay. As shown in Fig. 2a, SPAS-1115–172 interacted with tubulin and thus we identified this region MTBD. It is interesting to mention that in all spastin homologues from human, mouse, fly and C. elegans, MTBD is located between MIT and AAA domains, although their primary sequences are not well conserved (Fig. S1 in Supporting Information).
Modeled tertiary and quaternary structure of the AAA domain of SPAS-1
The complete AAA domain sequence of SPAS-1 was searched against PDB using SWISS-MODEL, and the Drosophila spastin monomer was found to be the closest homologue of SPAS-1 and show 50% sequence identity over the length of the AAA domain. Therefore, the tertiary structure model of AAA domain of SPAS-1 was created using homology modeling with Drosophila spastin monomer as a template (PDB code: 3b9p) (Roll-Mecak & Vale 2008). The closest homologue of known structure, which forms a hexamer, is D1 domain of p97/VCP (PDB code: 1r7r) (Huyton et al. 2003). We generated a hexameric structure of tertiary structure model of SPAS-1 AAA domain using p97/VCP as a template (Fig. 3a).
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The AAA domain contains the highly conserved aromatic residue that lies in the pore region of hexamer (Yamada-Inagawa et al. 2003; Schlieker et al. 2004; Gerega et al. 2005; Scott et al. 2005b). In the E. coli AAA protease FtsH, the conserved aromatic residue (F228) is important for substrate binding and subsequent translocation of substrates into the protease chamber (Yamada-Inagawa et al. 2003). C. elegans SPAS-1 has a tryptophan residue (W251) at the corresponding position to E. coli FtsH F228 (Fig. 3a). To elucidate the importance of the conserved aromatic residue in the pore motif, the tryptophan residue was replaced with alanine, glutamic acid, lysine or phenylalanine. As shown in Fig. 3b, all mutant proteins retained ATPase activity. We then examined effects of these mutations on microtubule severing. When wild-type SPAS-1 was expressed in cultured cells, microtubule network clearly disappeared (Fig. 3b), indicating that SPAS-1 possesses microtubule-severing activity as previously reported (Matsushita-Ishiodori et al. 2007). Although SPAS-1W251F as well as wild-type showed microtubule-severing activity, SPAS-1W251A, SPAS-1W251E and SPAS-1W251K did not show it. These data indicate that the conserved aromatic residue in the pore motif is required for microtubule severing.
Recognition of C-terminal amino acid residues of tubulin by the pore loop of SPAS-1
It has been reported that the pore motif of human spastin recognizes the C-terminal region of -tubulin, which is acidic and unstructured (Nogales et al. 1998, 1999; White et al. 2007). It should be noted that this recognition is independent of MTBD (White et al. 2007). To examine whether SPAS-1 recognizes the C-terminal acidic tail of -tubulin, basic amino acid residues exposed on surface based on the modeled SPAS-1 structure, R176, K205, K236, K257, R260, R267, R286, R295 and R296, were replaced by alanine (Fig. 4a,b). As clearly shown in Fig. 4c, when SPAS-1K257A, SPAS-1R260A, SPAS-1R286A, SPAS-1R295A and SPAS-1R296A mutants (pore mutants), whose mutated residues are locating in the pore region, were expressed, microtubule-severing activity was abolished. In contrast, in the case of non-pore mutants (SPAS-1R176A, SPAS-1K205A, SPAS-1K236A and SPAS-1R267A) on Face A, microtubule network disappeared (Fig. 4c). It is worth mentioning that K257, R260, R286, R295 and R296 are all conserved among human, mouse, fly and C. elegans spastin homologues (Fig. S1 in Supporting Information). These data suggest that the basic amino acid residues in the pore region play an important role in microtubule severing.
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-tubulin in vitro. Biosensor binding assays were performed by using SPAS-1 and -tubulin C-terminal 22 amino acid residues fused to glutathione S-transferase (GST). As shown in Fig. 5, response units increased proportionally with increasing concentrations of SPAS-1 with a dissociation constant of a KD of 0.03 µM, but not with bovine serum albumin (BSA), implying that the C-terminal region of -tubulin is recognized specifically by SPAS-1. To further demonstrate the direct interaction between SPAS-1 pore residues and the C-terminal region of -tubulin, we also performed the biosensor binding assays using pore mutants of SPAS-1, such as W251A and K257A. KD values of pore mutants were almost equal to that of wild-type (KD values of W251A and K257A were 0.03 and 0.04 µM respectively). These results suggest that there are multiple residues of pore region for binding to the C-terminal region of -tubulin.
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Discussion |
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It has been reported that AAA proteins generally assemble into a hexamer (Ogura & Wilkinson 2001; Yakushiji et al. 2006; White et al. 2007; Pantakani et al. 2008; Roll-Mecak & Vale 2008). In particular, katanin, Vps4p and spastin, which are classified into the same subgroup of AAA proteins, form a hexamer in an ATP-dependent manner (Babst et al. 1998; Hartman & Vale 1999; White et al. 2007; Roll-Mecak & Vale 2008). We demonstrated, however, that wild-type SPAS-1 and ATP-binding and -hydrolysis defective mutants SPAS-1K224R and SPAS-1E278Q formed a stable hexamer in a concentration-dependent manner (Fig. 1f). In contrast, SPAS-1 of low concentrations did not form a stable hexamer, rather it shows a broad elution profile over monomer to hexamer, even in the presence of ATP (Fig. 1g). We also demonstrated that MTBD of SPAS-1 is essential for binding of SPAS-1 to microtubules (Fig. 2). Furthermore, ATPase activity of SPAS-1 was greatly stimulated by the addition of microtubules (Fig. 1e). Taken these results together, we propose that MTBD of SPAS-1 plays a critical role in enrichment of SPAS-1 to microtubules, where SPAS-1 is concentrated and able to form a stable hexamer, subsequently its ATPase activity is stimulated.
We modeled tertiary and quaternary structures of the AAA domain of SPAS-1 based on template crystal structures of Drosophila spastin monomer and p97/VCP hexamer (Fig. 3a) (Huyton et al. 2003; Roll-Mecak & Vale 2008). The AAA domain contains the highly conserved aromatic residue that lies in the pore region of hexamer and plays a crucial role for function of AAA proteins (Yamada-Inagawa et al. 2003; Scott et al. 2005b; White et al. 2007; Roll-Mecak & Vale 2008). Our mutational analyses revealed that the SPAS-1W251F mutant still has microtubule-severing activity, while SPAS-1W251A, SPAS-1W251E and SPAS-1W251K lost it (Fig. 3b). These results clearly indicate that the conserved aromatic residue in the pore region of SPAS-1 is important for its microtubule-severing activity.
Interestingly, subtilisin treatment of microtubules, which removes small peptides from the C terminus of - and β-tubulin, renders microtubules resistant to severing by both spastin and katanin (McNally & Vale 1993; Roll-Mecak & Vale 2005). The C-terminal tail of tubulin is extremely acidic and unstructured (Nogales et al. 1998, 1999). It has been reported that the pore loops in hexameric spastin recognize the extreme C terminus of tubulin and that this recognition is critical for microtubule severing (White et al. 2007). Our mutational analyses on solvent exposed, basic amino acid residues revealed that the basic amino acid residues in the pore region, K257, R260, R286, R295 and R296, are important for microtubule severing (Fig. 4). It should be again noted that all of these amino acid residues are completely conserved among spastin homologues from human, mouse, fly and C. elegans. In contrast, when mutations were introduced on the solvent exposed, basic amino acid residues out of the pore region, microtubule-severing activity was not affected (Fig. 4). Furthermore, we detected the direct interaction of the C-terminal polypeptide of tubulin with SPAS-1 (Fig. 5). To the best of our knowledge, this is the first report of their direct interaction. These data strongly demonstrate that tubulin access to SPAS-1 from Face A side, in agreement with the previous report (Roll-Mecak & Vale 2008). Furthermore, single mutation of pore residues did not significantly affect the KD value between SPAS-1 and C-terminal region of -tubulin. These results suggest that there are multiple residues for binding to the C-terminal region of -tubulin.
As SPAS-1K257A mutant colocalized with microtubules similar to SPAS-1E278Q mutant, K257 is critical to dissociation from microtubules (Fig. 4c). It should be noted that SPAS-1K257A mutant possessed the ATPase activity comparable to that of wild-type SPAS-1 and was oligomerized normally. It has been reported that cycles of ATP hydrolysis induce conformational changes of the conserved aromatic residue in the pore region, which facilitates translocation of substrates (Yamada-Inagawa et al. 2003). Interestingly, K257 is located close to the corresponding aromatic residue W251. Taken together, it is likely that K257 is involved in not only recognition of microtubules but also an energy transduction event leading to conformational change. Intriguingly, we further demonstrated that additional mutations on W251, R260 and R296, which expose on Face A in the pore region (Fig. 4a,b), suppressed colocalization of SPAS-1 with microtubules due to the K257A mutation, while mutations on R286A and R295A, which locate inside the pore, did not (Fig. 6). These results imply that there are at least two recognition sites in the pore region, one is a set of basic amino acid residues on Face A and the other is a set of basic amino acid residues inside the pore. These results also indicate that the recognition on Face A is preceded to the recognition of inside the pore, that is, substrates may be translocated from Face A side to inside the pore. Similar results were obtained in a recent report as two-step model for ClpX (Martin et al. 2008).
Consequently, we propose two models of microtubule severing by SPAS-1, in which SPAS-1 adopts ‘dissociation’ or ‘translocation’ system (Fig. 7). SPAS-1 monomers interact with microtubules through MTBD of SPAS-1 and thus are enriched on microtubules, leading to form a hexamer in an ATP-independent manner. In the ‘dissociation’ model (Fig. 7a), the solvent exposed, basic amino acid residues in the pore first interact with the extremely acidic and unstructured C-terminal peptides of /β-tubulin, that is, SPAS-1 traps a tubulin at the pore region. Next, the conformational change of SPAS-1 oligomers resulting from cycles of ATP hydrolysis may give rise to disintegration of tubulin from microtubules and release of tubulin together with SPAS-1 oligomers. Subsequently, SPAS-1 oligomers may be dissociated due to the disappearance of anchorage. These tubulin extraction steps will be repeated at the region extracted previously or at random. Thus, microtubules will be destabilized and finally disassembled. Alternatively, dissociation of SPAS-1 oligomers might be occurred simultaneously with microtubule disassembly. This type of dissociation model has been demonstrated for katanin, NSF, and PEX1 and PEX6 actions (Hanson et al. 1997; Hartman & Vale 1999; Platta et al. 2005). The important property of this model is that dissociated substrates may not be completely unfolded and are ready to be recycled (McNally & Vale 1993; Hanson et al. 1997; Platta et al. 2005).
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/β-tubulin dimer through MTBD. This type of translocation model has been demonstrated for FtsH, ClpXP, ClpAP and 19S regulatory particle of proteasome (Voges et al. 1999; Kim et al. 2000; Ishikawa et al. 2001; Yamada-Inagawa et al. 2003). This system seems to relate to unfoldases coupled with proteases. However, this model has recently been suggested for spastin and Vps4p (Kieffer et al. 2008; Roll-Mecak & Vale 2008). The fundamental distinction of these two models is the direction of energy: in the dissociation model, an ATP-driven conformational change in SPAS-1 is relayed to dissociation of SPAS-1 hexamer, while in the translocation model, it is relayed to translocate the substrate tubulin. On the other hand, it has recently been reported that ClpB, which extracts unfolded polypeptides from aggregates via substrate translocation through its central pore, partially translocates the misfolded protein and subsequently dissociates from trapped substrates by instability of ClpB hexamer (Haslberger et al. 2008). Further experiments will be needed to generate a comprehensive picture of microtubule severing by spastin.
It is interesting to discuss that SPAS-1, particularly SPAS-1E278Q, may form a dodecamer, molecular mass of c. 820 kDa (Fig. 1f). It has been reported that Vps4p forms a dodecamer comprising two stacked hexameric rings and that the C-terminal -helix of the AAA domain plays a critical role in the formation of dodecamer (Babst et al. 1998; Scott et al. 2005b; Vajjhala et al. 2008). SPAS-1 also has a similar C-terminal -helix (Fig. S3 in Supporting Information). Therefore, it is possible that cellular activity of SPAS-1 may be regulated by the formation of hexamer/dodecamer. The dodecameric formation may be enhanced by ATP binding, as it was more prominent in SPAS-1E278Q, the ATP-hydrolysis deficient mutant (Fig. 1f). In the case of Vps4p, Vta1p has been identified to be involved in the formation and maintenance of dodecamer (Azmi et al. 2006; Lottridge et al. 2006; Vajjhala et al. 2006; Xiao et al. 2008). The interesting question whether SPAS-1 requires such a factor to form a dodecamer needs to be addressed.
The N-terminal region of AAA proteins is rich in diversity and considered to determine their substrate specificity. Spastin is composed of the MIT domain and MTBD in the N-terminal region (Fig. 2a) (Takasu et al. 2005; White et al. 2007). While MTBD is a fundamental domain for association with microtubules as discussed above, the MIT domain is not involved in association with microtubules (Fig. 2a) and microtubule severing (data not shown). It is interesting to mention that the ESCRT-III complex-associated endosomal protein CHMP1B has been identified as an interactor of spastin (Reid et al. 2005). The MIT domain of Vps4p has been recently verified to be a determinant for binding to ESCRT-III proteins (Scott et al. 2005a; Obita et al. 2007; Stuchell-Brereton et al. 2007; Vajjhala et al. 2007; Shim et al. 2008; Xiao et al. 2008). Therefore, it is reasonable to speculate that spastin homologues may also be involved in endosomal functions through the MIT domain.
Finally, we summarize effects of disease-associated mutations (K224R and R296A) on microtubule-severing activity. SPAS-1K224R mutant did not possess microtubule-severing activity (Matsushita-Ishiodori et al. 2007) due to the defective ATPase activity (Fig. 1d). The microtubule-severing activity of SPAS-1 was also impaired by the R296A mutation (Fig. 4c), which cause the defective tubulin recognition in the pore region as discussed above. It is thus possible that the disease-associated mutations prevent the microtubule-severing activity of spastin and cause pathological effects. Future studies will be required to reveal the detailed mechanisms of microtubule severing and HSP.
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Experimental procedures |
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Cell culture, transfection and immunocytochemistry were performed as described previously (Matsushita-Ishiodori et al. 2007).
Plasmid construction
Full-length cDNA fragment for spas-1 was amplified with primers 5'-ATCACATATGTTCGCCTTTTCAAAAGG-3' (sense, NdeI site underlined) and 5'-AGAACCCGGGTTAGCAACCGAAACTTCGAG-3' (antisense, SmaI site underlined) from yk735e10 (a generous gift from Dr Y. Kohara, National Institute of Genetics, Japan), and cloned into pET-15b, yielding pCKX1230. Site-directed mutagenesis in spas-1 was carried out using QuikChange II XL Site-directed mutagenesis kit (Stratagene, Cedar Creek, TX). The mutations are as follows: K224R (AAA to AGA), E278Q (GAA to CAA), W251A (TGG to GCG), W251K (TGG to AAG), W251E (TGG to GAG), W251F (TGG to TTT), K257A (AAA to GCA), K257R (AAA to AGA), K257Q (AAA to CAA), K257E (AAA to GAA), R260A (CGA to GCA), R286A (CGA to GCA), R295A (AGA to GCA), R296A (AGA to GCA), R176A (CGG to GCG), K205A (AAA to GCA), K236A (AAG to GCG), R267A (CGA to GCA) and C284Y (TGT to TAT). For transfection experiments, entire open reading frames encoding wild-type and mutant SPAS-1 proteins were amplified and cloned into the modified pcDNA3, from which an N-terminal FLAG-epitope tagged fusion protein is expressed. DNA segment encoding C-terminal 22 amino acids of C. elegans-tubulin (KDYEEVGADSNEGGNEEEGEEY) was amplified with primers 5'-GCCCGGATCCATGAAGGACTACGAAGAGGTCGG-3' (sense, BamHI site underlined) and 5'-GCCCGCGGCCGCTTTAATACTCTTCTCCTTCC-3' (antisense, NotI site underlined) from yk1300d1 (a generous gift from Dr Y. Kohara), and cloned into pGEX-6P-3, yielding pCKX6136. Plasmid DNAs were column purified (Qiagen, Valencia, CA). All of the mutations were confirmed by sequencing.
Purification of proteins
SPAS-1 plasmids described above were introduced into E. coli BL21(DE3). Transformants were grown at 30 °C, and expression of wild-type and mutant SPAS-1 was induced by the addition of isopropyl-β-D-thiogalactopyranoside (IPTG) (0.5 mM), followed by growth for 3 h. Cells from 1.5 L culture were harvested and suspended in 22.5 mL of lysis buffer [50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 0.2% NP-40, 20 mM imidazole, 1 M arginine hydrochloride and 0.1 mM ATP] supplemented with complete protease inhibitor cocktail (Nacalai Tesque, Kyoto, Japan). After sonication on ice, cell lysates were ultracentrifuged at 100 000 x g for 1 h at 4 °C, and the supernatant was dialyzed against buffer A [50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 0.2% NP-40, 20 mM imidazole and 0.1 mM ATP]. Soluble fractions thus obtained were incubated with Ni2+-NTA agarose (Qiagen) for 1 h at 4 °C. His-tagged recombinant SPAS-1 proteins were finally eluted with Elution buffer [50 mM Tris–HCl (pH 7.5), 50 mM NaCl, 0.1% NP-40, 0.1 mM ATP, 10% glycerol and 200 mM imidazole]. Eluted fractions were analyzed by SDS–PAGE and visualized with CBB (Coomassie brilliant blue) staining. Proteins were concentrated using Amicon Ultra-15 (Millipore, Billerica, MA), dialyzed against storage buffer [25 mM Tris–HCl (pH 7.5), 50 mM NaCl, 0.01% NP-40 and 10% glycerol], and stored at –80 °C. For CD spectrum and size exclusion column chromatography, NP-40 was omitted from elution and storage buffers.
GST-fusion proteins were expressed in E. coli XL-1 Blue. Transformants were grown at 37 °C, and expression of GST-fusion proteins was induced by the addition of IPTG (1 mM), followed by growth for 3 h. Cells from 100 mL culture were harvested and resuspended in phosphate-buffered saline buffer containing 1 mg/mL lysozyme. After lysed with 0.2% Triton X-100, soluble fractions were incubated with Glutathione Sepharose 4B (GE Healthcare, Little Chalfont, UK) for 30 min at room temperature. GST-fusion proteins were eluted with 50 mM Tris–HCl (pH 8.0) and 10 mM reduced glutathione. Samples were analyzed by SDS–PAGE as described above. Eluted fractions were dialyzed against storage buffer [50 mM Tris–HCl (pH 8.0), 20 mM NaCl, 10 mM mercaptoethanol and 10% glycerol]. Purified GST-fusion proteins were stored at –80 °C. Protein concentration was determined by SDS–PAGE, followed by densitometric scanning of CBB stained gels. BSA was used as a standard.
Malachite ATPase assay
ATPase activity was measured at 30 °C using the malachite green colorimetric assay as described previously (Akiyama et al. 1996). Sodium phosphate was used as a standard.
Microtubules were prepared by incubating bovine brain /β-tubulin dimers (Cytoskeleton) (5 mg/mL) at 37 °C for 45 min in PEM buffer (160 mM PIPES [pH 6.9], 4 mM MgCl2, 1 mM EGTA) with 1 mM GTP. Taxol was then added to a final concentration of 20 µM. After 10 min at 37 °C, microtubules were separated from soluble tubulin by ultracentrifugation. Taxol-stabilized microtubules were used for ATPase assay.
Size exclusion column chromatography
The oligomeric state of SPAS-1 was analyzed by size exclusion column chromatography using a Superose 6 10/300GL column (GE Healthcare). Samples were applied to the column in a buffer consisting of 25 mM Tris–HCl (pH 7.5), 500 mM NaCl and 10% glycerol. Proteins eluted were monitored by absorbance at 280 nm and analyzed by SDS–PAGE.
In vitro pull-down assay
For pull-down assays, 20 µg His-tagged proteins coupled to Ni2+-NTA agarose (Qiagen) (75 µL) were incubated in 50 mM Tris–HCl (pH 7.5), 0.1% Triton X-100, complete protease inhibitor cocktail (Nacalai Tesque) and 3 µg bovine brain /β-tubulin dimers (Cytoskeleton) under continuous shaking for 3 h at 4 °C. After washing, bound proteins were eluted with 200 mM imidazole and analyzed by SDS–PAGE followed by immunoblotting using anti--tubulin antibody.
Surface plasmon resonance detection
The surface plasmon resonance experiments were performed with Biacore 2000 instrument (GE Healthcare). CM4 sensor chips and the Amine Coupling Kit (N-hydroxysuccinimide, N-ethyl-N'-(3-diethyl-aminopropyl)-carbodiimide, ethanolamine-hypochloride) were from GE Healthcare. To analyze the binding of SPAS-11–152 to tubulin, the first flow cell of CM4 sensor chip was coupled with no protein to be used as a reference flow cell, and the second flow cell was coupled with /β-tubulin dimer. The coupling was performed in 10 mM sodium acetate (pH 4.4) using Amine Coupling Kit. To detect the recognition of the C-terminal peptides of -tubulin by SPAS-1, the first flow cell of CM4 sensor chip was coupled with GST-fusion protein to be used as a reference flow cell, and the second flow cell was coupled with GST-fusion proteins representing the C-terminal 22 amino acids of -tubulin. The coupling was performed in 10 mM sodium acetate (pH 5.0) using Amine Coupling Kit. All binding experiments were performed in 10 mM Hepes (pH 7.6), 150 mM NaCl, 0.005% Tween20, 1 mM DTT, 1 mM MgCl2 and 3 mM ATP. The respective SPAS-1 proteins were applied to the sensor chip at the flow rate of 20 µL/min for 2 min, and temperature was maintained at 25 °C. The response curve on sample was subtracted from reference flow cell.
Modeling of tertiary and quaternary structures of the AAA domain of SPAS-1
The complete AAA domain sequence of SPAS-1 was searched against PDB using the SWISS-MODEL program (http://swissmodel.expasy.org), with default parameters, to identify closely related homologues of SPAS-1 with known 3D structure. The AAA ATPase domain of SPAS-1 was modeled on the basis of the tertiary structures of the template (PDB code: 3b9p) (Roll-Mecak & Vale 2008). The copies of modeled tertiary structure were assembled to form a hexameric quaternary assembly on the basis of the hexameric template (PDB code: 1r7r) (Huyton et al. 2003). This modeled quaternary structure was energy minimized using the Swiss PDB viewer (http://spdbv.vital-it.ch/).
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Footnotes |
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* yamanaka{at}gpo.kumamoto-u.ac.jp or ogura{at}gpo.kumamoto-u.ac.jp |
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Received: 15 December 2008
Accepted: 2 May 2009
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) at 222 nm. The melting temperature was calculated as 47 °C from the reflection points. (d) ATPase activities of wild-type and mutant SPAS-1 (K224R and E278Q) were measured at 37 °C. (e) ATPase activities of wild-type SPAS-1 were measured in the presence of microtubules at 37 °C. ATPase activities are plotted with increasing amounts of substrates. (f) Oligomer formation of SPAS-1 at a high concentration. Wild-type (5 mg/mL) (solid line) and mutant (K224R, 10 mg/mL, and E278Q, 5 mg/mL) SPAS-1 proteins (dashed line and broken line respectively) were fractionated on a Superose 6 10/300GL column as described under Experimental procedures. It should be mentioned that the buffer used for chromatography did not contain ATP. Proteins eluted were monitored by absorbance at 280 nm. Molecular mass markers: void volume, V0; thyroglobulin, 669 kDa; ferritin, 440 kDa; catalase, 232 kDa; BSA, 67 kDa. (g) Oligomer formation of SPAS-1 at a low concentration. Wild-type (0.55 mg/mL) and SPAS-11–210 (0.55 mg/mL) were fractionated on a Superose 6 10/300GL column as described under Experimental procedures. Proteins eluted were analyzed by SDS–PAGE and visualized by silver staining. Molecular mass markers: catalase, 232 kDa; BSA, 67 kDa; chymotrypsinogen A, 25 kDa.
