Physical and functional connection between auxilin and dynamin during endocytosis
2006; Springer Nature; Volume: 25; Issue: 18 Linguagem: Inglês
10.1038/sj.emboj.7601298
ISSN1460-2075
AutoresSanja Sever, Jesse Skoch, Sherri L. Newmyer, Rajesh Ramachandran, David Ko, Mary McKee, Richard Bouley, Dennis A. Ausiello, Bradley T. Hyman, Brian J. Bacskai,
Tópico(s)Microtubule and mitosis dynamics
ResumoArticle31 August 2006free access Physical and functional connection between auxilin and dynamin during endocytosis Sanja Sever Corresponding Author Sanja Sever Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Jesse Skoch Jesse Skoch Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Sherri Newmyer Sherri Newmyer GW Hooper Foundation, The University of California, San Francisco, CA, USA Search for more papers by this author Rajesh Ramachandran Rajesh Ramachandran Department of Cell Biology, The Scripps Research Institute, La Jolla, CA, USA Search for more papers by this author David Ko David Ko Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Mary McKee Mary McKee Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Richard Bouley Richard Bouley Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Dennis Ausiello Dennis Ausiello Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Bradley T Hyman Bradley T Hyman Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Brian J Bacskai Brian J Bacskai Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Sanja Sever Corresponding Author Sanja Sever Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Jesse Skoch Jesse Skoch Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Sherri Newmyer Sherri Newmyer GW Hooper Foundation, The University of California, San Francisco, CA, USA Search for more papers by this author Rajesh Ramachandran Rajesh Ramachandran Department of Cell Biology, The Scripps Research Institute, La Jolla, CA, USA Search for more papers by this author David Ko David Ko Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Mary McKee Mary McKee Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Richard Bouley Richard Bouley Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Dennis Ausiello Dennis Ausiello Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Bradley T Hyman Bradley T Hyman Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Brian J Bacskai Brian J Bacskai Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA Search for more papers by this author Author Information Sanja Sever 1, Jesse Skoch2, Sherri Newmyer3, Rajesh Ramachandran4, David Ko1, Mary McKee1, Richard Bouley1, Dennis Ausiello1, Bradley T Hyman2 and Brian J Bacskai2 1Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA 2Alzheimer's Disease Research Laboratory, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA, USA 3GW Hooper Foundation, The University of California, San Francisco, CA, USA 4Department of Cell Biology, The Scripps Research Institute, La Jolla, CA, USA *Corresponding author. Renal Unit, Nephrology Division, Department of Medicine, Harvard Medical School and Massachusetts General Hospital, Charlestown, MA 02129, USA. Tel.: +1 617 724 8922; Fax: +1 617 726 5669; E-mail: [email protected] The EMBO Journal (2006)25:4163-4174https://doi.org/10.1038/sj.emboj.7601298 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info During clathrin-mediated endocytosis, the GTPase dynamin promotes formation of clathrin-coated vesicles, but its mode of action is unresolved. We provide evidence that a switch in three functional states of dynamin (dimers, tetramers, rings/spirals) coordinates its GTPase cycle. Dimers exhibit negative cooperativity whereas tetramers exhibit positive cooperativity with respect to GTP. Our study identifies tetramers as the kinetically most stable GTP-bound conformation of dynamin, which is required to promote further assembly into higher order structures such as rings or spirals. In addition, using fluorescence lifetime imaging microscopy, we show that interactions between dynamin and auxilin in cells are GTP-, endocytosis- and tetramer-dependent. Furthermore, we show that the cochaperone activity of auxilin is required for constriction of clathrin-coated pits, the same early step in endocytosis known to be regulated by the lifetime of dynamin:GTP. Together, our findings support the model that the GTP-bound conformation of dynamin tetramers stimulates formation of constricted coated pits at the plasma membrane by regulating the chaperone activity of hsc70/auxilin. Introduction Clathrin-mediated endocytosis is a multistep process that involves formation of clathrin-coated vesicles (CCVs) from the plasma membrane (Kirchhausen, 2000). Based on current models, coat assembly starts with the recruitment and oligomerization of adaptor proteins and is followed by the recruitment of clathrin. Early stages of invagination are followed by a well-defined constriction step that requires the GTPase dynamin, whose exact role in this process is controversial. Based on the classical model, self-assembly of dynamin into higher order structures such as rings and spirals is thought to drive the constriction step, whereas assembly-stimulated GTPase activity is thought to be directly involved in severing the neck of a coated pit (for a review see Sever et al, 2000b). Alternatively, we have proposed that dynamin functions as a regulatory GTPase whose GTP-bound form regulates coat rearrangement and thereby drives constriction. In this view, assembly-stimulated GTP hydrolysis acts as a 'switch off' mechanism for dynamin's active form (Sever et al, 1999; Narayanan et al, 2005). After vesicle release, the clathrin coat is disassembled by hsc70 (Schlossman et al, 1984) and its cochaperones, aux1 (neuronal isoform) (Ungewickell et al, 1995) or auxilin-2 (aux2) (ubiquitous form) (Greener et al, 2000; Umeda et al, 2000). Although the N-terminus of aux2 is a functional protein kinase that is absent from auxillin-1, (aux1) the two proteins are otherwise highly homologous, sharing three key domains: the tensin domain, a clathrin-binding domain and a C-terminal DnaJ domain that interacts with hsc70. Both proteins can induce clathrin polymerization into baskets and they both support uncoating of CCVs by hsc70 in vitro (Ungewickell et al, 1995; Greener et al, 2000; Umeda et al, 2000). The role of hsc70/auxilin in endocytosis does not appear to be restricted to the uncoating reaction. Dominant-negative hsc70 mutants have been shown to inhibit the formation of constricted coated pits (Newmyer and Schmid, 2001), the same step that requires dynamin. Furthermore, experiments in Drosophila identified genetic interactions between dynamin and hsc70 (Chang et al, 2002), and between dynamin and the DnaJ domain cochaperone of hsc70, Rme-8 (Chang et al, 2004), suggesting that dynamin and chaperones function in the same pathway in vivo. Further strengthening the connection between chaperones and dynamin, we have identified direct interactions between the GTP-bound form of dynamin and aux1, as well as hsc70 (Newmyer et al, 2003). Based on these results, we proposed that dynamin: GTP regulates hsc70/auxilin to drive CCV formation. This model was recently supported by two studies showing that downregulation of aux2 in HeLa cells inhibits clathrin-mediated endocytosis, but they differed with regard to the steps that required aux2. Downregulation of aux2 by shRNA resulted in the overall loss of clathrin-coated pits, suggesting a role for aux2 in de novo pit formation (Lee et al, 2005), whereas experiments performed using siRNA place the role of aux2, and specifically its cochaperone activity, somewhere between the initiation and the uncoating steps (Zhang et al, 2005). Live-cell imaging of clathrin (Merrifield et al, 2005) and downregulation of clathrin in cells (Iversen et al, 2003) together suggested that clathrin assembly may play a role in the final stages of invagination that lead to membrane scission. As there is no evidence that auxilin and hsc70 remove clathrin from clathrin-coated pits at the plasma membrane, but only from vesicles after budding (Ungewickell et al, 1995), our data suggest that interactions between dynamin and the chaperone machinery might suppress the uncoating activity. Here we pinpoint constriction of clathrin-coated pits as the step in endocytosis that requires the cochaperone activity of auxilin. The identical step was previously shown to be regulated by the lifetime of dynamin:GTP (Sever et al, 2000a; Narayanan et al, 2005), consistent with auxilin being a bona fide dynamin downstream effector. Supporting this idea, fluorescence lifetime imaging microscopy (FLIM) shows that interactions between dynamin and auxilin are GTP- and endocytosis-dependent. In addition, we show that in order to interact with auxilin in vivo, dynamin must self-associate into tetramers. Together, the data support the model that dynamin tetramers regulate constriction of clathrin-coated pits by direct interactions with hsc70/auxilin. Results Endogenous dynamin-2 and aux2 occupy the same coated profiles Our previous biochemical experiments showed that hsc70/auxilin interacts specifically with the GTP-bound form of dynamin, suggesting that this chaperone could be a dynamin effector. To further address this possibility, we asked whether auxilin and dynamin occupy the same coated profiles in vivo. We used the kidney cortex as a model system. The apical membranes of kidney proximal tubules support extensive clathrin-mediated endocytosis to retrieve proteins from the urine, and thus contain high levels of endocytic proteins (Sun et al, 2002). As shown in Figure 1A, left panel, anti-aux1 antibodies raised against auxilin405−814 specifically recognized auxilin-1 (aux1) in rat brain cytosol (RBC), and they also specifically recognized aux2 in inner medullary collecting duct (IMCD) cell extracts. Staining of kidney proximal tubules for aux2 showed that this protein is present in the apical membrane (Figure 1B, full arrow), as well as at the basal membrane, but at significantly lower levels (Figure 1B, open arrow). The same staining pattern was exhibited by dynamin (Figure 1B') and clathrin (data not shown). Because immunofluorescence cannot distinguish between auxilin on CCVs versus auxilin on coated pits, we performed immunogold electron microscopy. Kidney cortex slices were incubated with polyclonal anti-aux1 antibodies coupled to 15 nm gold particles (Figure 1C). Aux2 antigenic sites were concentrated on invaginated regions that could be readily identified as clathrin-coated pits on the basis of the thick, electron-dense coat of clathrin that characterizes these structures in electron micorgraphs (Figure 1C; Sun et al, 2002). Most coated profiles are connected with the membrane and thus do not represent budded vesicles. We next used monoclonal anti-dynamin antibodies in addition to polyclonal anti-aux1 antibodies (Figure 1D, E and F). In this case, aux2 (large gold particles) and dynamin-2 (dyn2) (small gold particles) antigenic sites were present on the same coated profiles (Figure 1D, E and F). In some instances, dynamin is found concentrated at the neck of invaginated coated pits (Figure 1E, arrow). It is important to note that dynamin is not present on CCVs after they have budded (Damke et al, 1994; and our data not shown). Therefore, the presence of aux2 on the same coated profiles as dyn2 indicates that it localizes to coated pits before budding. Figure 1.Endogenous aux2 and dyn2 colocalize at the same clathrin-coated profiles. (A) Aux1 antibodies recognize aux2. Western blot analysis of 100 μg cytosol prepared from IMCD cells transiently transfected with aux2 (lane 1), untransfected IMCD cells (lane 2), RBC (lane 3), and 0.1 μg purified recombinant aux1 (lane 4). The same samples were also blotted with anti-aux2 antibodies (Stressgene) (right panel). (B) Aux2 and dyn2 localize to the same apical region of the kidney proximal tubules. Rat kidneys were stained with aux1 antibodies (left panel) or hudy-1 (right panel). Closed arrow, apical membrane; open arrow, basal membrane of the polarized epithelial cells that form proximal kidney tubules. (C, D, E, F) Immunogold electron microscopy showing aux2 antigenic sites concentrated on clathrin-coated pits at the apical plasma membrane. Kidney cortex was labeled with only auxilin (15 nm gold) (C), or both auxilin (15 nm gold) and dynamin (10 nm gold) antibodies (D, E, F). The scale bars correspond to 0.1 μm. Download figure Download PowerPoint Detection of GTP-dependent dynamin–auxilin interactions in cells using FLIM To examine whether dynamin directly interacts with auxilin in vivo, we performed FLIM. The fluorescence lifetime of a high-energy donor fluorophore is influenced by its surrounding microenvironment, and it is shortened in the immediate vicinity of a lower energy acceptor fluorophore. Thus, detection of shortened lifetimes demonstrates fluorescence resonance energy transfer (FRET) and indicates spatial proximity of the two labeled molecules. We first sought a cell type where adenoviral expression of wild-type aux1 (auxWT) has no negative effect on endocytosis. Overexpression of auxWT in IMCD cells did not cause accumulation of clathrin into cytosolic granules (Supplementary Figure 1A and C and compare with Supplementary Figure 3A) or inhibit endocytosis of rhodamine-transferrin (R-Tfn) (Figure 5C). Confocal microscopy of IMCD cells coinfected with adenoviruses expressing dyn1 and aux1 detected a similar distribution of both proteins in the cytoplasm, and enrichment at the edge of the cellular monolayer (Figure 2A, white arrows). A similar enrichment at the edge of the cellular monolayer was observed for clathrin but only after expression of auxWT (Supplementary Figure 1B), suggesting that this region supports active endocytosis. Association of the auxWT and dynWT with the membrane was further examined by subcellular fractionation. As shown in Figure 2B and C, endogenous dynamin was approximately equally distributed between the particulate (membrane bound) and soluble (cytoplasm) fractions, whereas endogenous auxilin was present predominantly in the cytoplasm (compare lanes 1 and 2). Overexpression of dynWT and auxWT led to an overall increase of these proteins in membrane-associated and soluble fractions (Figure 2), as previously observed for dynamin (van der Bliek et al, 1993). Although the exact mechanisms that target either dynamin or auxilin to the plasma membrane are not known (although binding of lipids (Lin et al, 1997) and clathrin-coated pits (Figure 1D; and Ungewickell et al, 1995) is likely involved), these data suggest that membrane targeting can be increased by overexpressing proteins. Figure 2.Identification of GTP- and endocytosis-dependent dynamin–auxilin interactions in IMCD cells. (A) Auxilin and dynamin colocalize at the edge of the cellular monolayer. IMCD cells were infected with adenoviruses encoding auxWT, dynWT or dynK44A, as indicated. Cells were optionally treated with MBCD before fixation. Cells were stained with polyclonal anti-auxilin antibodies followed by secondary IgG conjugated with the donor fluorophore a488, and monoclonal anti-dynamin antibodies hudy-1 followed by secondary IgG conjugated with acceptor fluorophore (a568. (B, C) Subcellular distribution of overexpressed dynamin and auxilin in IMCD cells. Cells were lysed and resolved into particulate and soluble fractions by centrifugation at 100 000 g. Total protein from both fractions was immunoblotted with hudy-1 (B) or anti-auxilin (C) antibodies. S, supernatant; P, pellet. (D) IMCD cells were infected and stained as described in (A). Intensity column shows auxilin distribution (a488). FLIM-ing column shows FRET between the donor and acceptor measured by the fluorescence lifetime, t1, of the donor fluorophore (a488) that is represented by a pseudo-colored image. (E) Lifetimes of the donor fluorophore (a488) under different experimental conditions. Each data point represents at least 10 images similar to those shown in (D). Download figure Download PowerPoint Cells expressing auxWT and dynWT were first stained with rabbit anti-auxilin antibodies followed by goat-anti-rabbit (GAR) antibody conjugated to Alexa 488 (a488), the donor fluorophore. We measured changes in the lifetime of a488 under different experimental conditions. In the absence of an acceptor fluorophore, the lifetime of a488 was 2228±53 ps (Table I). If the cells were also stained with goat anti-mouse secondary antibodies conjugated with a568 (GAMa568), the lifetime remained almost unchanged at ∼2188±62 ps. In contrast, staining cells with dog anti-goat secondary antibodies conjugated with a568 (DAGa568) as a positive control, resulted in a very short lifetime of 972±153 ps. Cells stained with both anti-auxilin (a488) and anti-dynamin antibodies (a568) resulted in a statistically significant reduction of a488 fluorescence lifetime to 1922±131 ps. The fastest rates of a488 lifetime were situated at the edges of the cellular monolayer (τ1=1722±134 ps; Figure 2D, FLIM-ing panel). This spatially restricted lifetime shortening in cells suggests that dynamin–auxilin interactions occur at the plasma membrane at the edges of the cellular monolayer. Table 1. FLIM analysis for proximity between auxilin and dynamin in IMCD cells Donor Acceptor Mean lifetime (τ1 in ps) (mean ±s.d.) (whole field) Lifetime (τ1 in ps) (mean ±s.d.) (fast edge-RDA) Mean Lifetime (whole field) P compared to Aux (a488) AuxWT (a488) None 2228±53 AuxWT (a488) GAM (a568) 2188±62 P>0.1 AuxWT (a488) DAG (a568) 972±153 P<0.05 AuxWT (a488) DynWT (a568) 1922±131 1722±134 P 0.1 AuxWT (a488) DynI690K (a568) 2186±39 None P>0.1 AuxWT (a488) DynK694A (a568) 1805±34 1605±187 P<0.05 AuxH875Q (a488) DynWT (a568) 1799±166 1708±136 P 0.1 Experiments were performed by staining cells with polyclonal anti-auxilin antibodies followed by goat anti-rabbit (GAR) secondary antibodies conjugated to Alexa-488 (a488, donor fluorophore), followed by monoclonal anti-dynamin antibody (hudy 1), followed by goat anti-mouse (GAM) secondary antibodies conjugated to Alexa-568 (a568, acceptor fluorophore). For positive control, cells were stained with donkey anti-goat (DAG) antibodies conjugated to Alexa-568 that recognize GAR-a488. τ2 was fixed to mean of Alexa-488 anti-auxilin antibody of 2207 picoseconds (ps). Range-dependent analysis (RDA) was performed by comparing only fast lifetimes at the edge of the cellular monolayer to the mean lifetime of 2207 ps, and it was determined as described in Materials and methods. For statistical analysis of residuals, one-way analysis of variance followed by a Tamhane post hoc test was performed with SPSS software. A P value of less than 0.05 constituted significance. Aux, auxilin; Dyn, dyanamin; IMCD, inner medullary collecting duct; MBCD, β-methyl-cyclodextrin. To test whether dynamin–auxilin interactions were dependent on clathrin-mediated endocytosis, we treated cells with β-methyl-cyclodextrin (MBCD), a chemical that depletes cholesterol from the plasma membrane and inhibits endocytosis at the stage of open coated pits that fail to deeply invaginate (Subtil et al, 1999). MBCD did not diminish dynamin–auxilin colocalization on the edges of the cellular monolayer (Figure 2A), or their membrane association (in Figure 2B and 2C, lane 6), but it abolished the FLIM signal (Figure 2D and 2E). This result suggests that dynamin–auxilin interactions might occur among the subset of these proteins that are associated with clathrin-coated pits, and that these interactions require a certain degree of coat curvature. It is important to note that whereas short lifetime only occurs if fluorophores are within a distance well below optical resolution, the pixels that are pseudocolored to reflect the presence or absence of short lifetime species are large compared to the distances involved for FRET. Next, we tested whether the observed dynamin–auxilin interactions are GTP-dependent. Cells were infected with adenoviruses expressing auxWT and dynK44A, a mutant of dynamin that cannot bind GTP. As shown in Figure 2D, there was no detectable FRET measured by FLIM. As dynamin targeting is not nucleotide dependent (Figure 2A and B; Song et al, 2004), loss of the signal cannot be attributed to an absence of dynK44A at the plasma membrane. The GTP dependence for dynamin–auxilin interactions in vivo is in agreement with in vitro binding experiments (Newmyer et al, 2003). Furthermore, these interactions were not dependent on a functional DnaJ domain, because an auxilin mutant with impaired J-domain function, auxH875Q, exhibited the same level of FRET as observed for auxWT (Figure 2E). This result is in agreement with our work that showed that auxilin fragments that lack the DnaJ domain, aux405−591 and aux591−814, both directly bind dynamin in vitro and potently inhibit endocytosis (Newmyer et al, 2003). Next, we examined whether dynK694A, a mutant of dynamin that increases the rate of endocytosis owing to impaired assembly into higher order structures (Sever et al, 1999), can interact with auxilin. DynK694A exhibited FRET as measured by FLIM (Figure 2E). The decrease in the lifetime of a488 in cells expressing dynK694A (τ1=1605±187 ps) is greater than in cells expressing dynWT (τ1=1722±134 ps), suggesting that dynK694A interacts with auxilin more efficiently than dynWT. These data support our original interpretation (Sever et al, 1999, 2000a) that dynK694A increases the rate of endocytosis because of improved interactions with effector proteins. A recent study showed that dynI690K is assembly incompetent and inactive for endocytosis (Song et al, 2004), and therefore concluded that dynamin self-assembly is essential for endocytosis. In our experiments, dynI690K exhibited no FRET as measured by FLIM (Figure 2E). The inability of this mutant to interact with auxilin could not be explained by defective targeting of dynI690K (Figure 2B, lane 10; Supplementary Figure 1D and Song et al, 2004). Interestingly, whereas dynI690K and dynK694A are both deficient in self-assembly, the former inhibits endocytosis whereas the latter is stimulatory. Our results therefore suggest that dynI690K inhibits endocytosis not due to a failure in self-assembly, but rather due to deficient interactions with auxilin. Dynamin exhibits positive cooperativity for GTP binding Although the K694A and I690K mutations are both situated within the GAP domain, which also functions as an assembly domain, dynI690K and dynK694A exhibit different biochemical properties; dynI690K is completely unable to assemble into higher order structures, even on lipids and microtubules (MT) (Song et al, 2004), whereas dynK694A exhibits a six-fold lower propensity to self-assemble, which is overcome by lipids or MT (Sever et al, 1999). Regardless of this difference, the FLIM results were puzzling as it was not obvious why differences in self-assembly should influence the interaction with auxilin. We therefore speculated that dynI690K was impaired not only for self-assembly, but also for GTP binding. This idea was inspired by the dynamin family member Dnm1, whose GTP binding is assembly dependent (Ingerman et al, 2005); to test it, we examined the interaction of dynI690K and dynK694A with GTP. GTP binding of dynamin was studied using a 'coupled GTPase assay' (Ingerman et al, 2005), in which GTP is continuously regenerated from guanosine 5′-diphosphate. Measurement of the GTPase activity of dynWT and dynK694A at high ionic strength (150 mM NaCl) revealed that the maximal rate of GTP hydrolysis for both proteins was ∼2.5 min−1 (Figure 3A and Table II). These data are similar to values determined using fixed time-point assays (Sever et al, 1999; Narayanan et al, 2005). Addition of the isolated GAP domain of dynamin, which mimics self-assembly into higher order structures (Sever et al, 1999), potently increased the rate of GTP hydrolysis by dynWT (Vmax=77 min−1), whereas it only partially increased the rate of GTP hydrolysis by dynK694A (Vmax=29 min−1, Figure 3B), in agreement with previous studies (Sever et al, 1999). In contrast to dynWT or dynK694A, dynI690K exhibited a three-fold lower basal rate of GTP hydrolysis, which could not be further stimulated by the addition of the GAP domain (Vmax=1.1 min−1, Figure 3A and B). All three proteins exhibited similar affinities for GTP under different conditions, in agreement with previously published data (Table II and Warnock et al, 1996). Importantly, the coupled assay revealed that dynWT and dynK694A both exhibit positive cooperativity with respect to GTP binding (Hill coefficient of ∼2, Supplementary Figure 2). This positive cooperativity was lost when dynamin self-assembly was promoted in low ionic strength, or through addition of the GAP domain (Hill coefficient=1, Table II). In contrast, dynI690K exhibited negative cooperativity with respect to GTP binding under all three experimental conditions (Hill coefficient=0.6–0.8, Supplementary Figure 2). Thus, the I690K mutation impaired the enzyme's ability to efficiently bind GTP, whereas K694A did not have this effect. Figure 3.Assembly-dependent GTP binding by dynamin. Steady-state kinetics of dynWT, dynK694A and dynI690K at 150 mM NaCl (A), or with 3 μM GAP domain at 50 mM NaCl (B). 0.06 mg/ml (0.6 μM) dynamin was assayed for GTPase activity. The difference between rates in three different experiments was less than 5%. (C) Analytical ultracentrifugation of 5 μM dynWT or dynI690K. (D, E) GTPase assays were initiated by the dilution of dynWT (D) or dynI690K (E) from 200 into 40 mM NaCl in the presence of 100 μM of GTP, and GTPase activity was measured over time. Download figure Download PowerPoint Table 2. Kinetic parameters of dynamin wild-type and assembly mutants Enzymes Vmax (min−1) Km (μM) Hill coefficient DynWT −High salt 2.3±0.2 40±2 1.9 −Low salt 4.8±0.5 35±4 1 +GAP 77±8 31±2 1 DynK694A −High salt 2.1±0.1 44±4 2.1 −Low salt 3.1±0.2 25±5 1 +GAP 29±5 33±2 1 DynI690K −High salt 0.7±0.1 25±5 0.6 −Low salt 0.8±0.1 25±4 0.7 +GAP 1.1±0.1 28±3 0.8 Dyn, dynamin. In its basal state at high ionic strength, dynamin is best described by a monomer–tetramer or dimer–tetramer equilibrium (Muhlberg et al, 1997; Binns et al, 1999). Differences in cooperativity of GTP binding between dynWT and dynI690K suggested that the I690K mutation might have changed the oligomeric state of dynI690K. As shown in Figure 3C, analytical ultracentrifugation revealed that in contrast to dynWT, which formed homotetramers, dynI690K was a homodimer. Thus, the I690K mutation abolished the ability of dynI690K to form tetramers, the first level of higher order structure. Together, the biochemical analysis suggests that to from a kinetically stable GTP-bound conformation, dynamin dimers need to oligomerize into a tetrameric state. In light of dynamin's oligomerization-dependent GTP-binding properties, we interpret the inability of dynI690K to bind auxilin as determined by FLIM as a consequence of its inability to form a kinetically stable GTP-bound conformation. Monitoring the rates of GTP hydrolysis continuously revealed that tetramer formation might be a prerequisite to assemble into higher order structures such as rings in solution. Dilution of dynamin from a high ionic strength buffer (tetramers) into a low ionic strength condition induces formation of higher order structures such as rings in solution (Hinshaw and Schmid, 1995), and leads to 2–3-fold increase in the rate of GTP hydrolysis by dynamin (Table II and Warnock et al, 1996). As shown in Figure 3D, when dynWT was diluted into low salt, t
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