Article8 June 2006free access Mechanism of endophilin N-BAR domain-mediated membrane curvature Jennifer L Gallop Jennifer L Gallop MRC Laboratory of Molecular Biology, Cambridge, UKPresent address: Department of Systems Biology, Harvard Medical School, Boston, MA 02115, USA Search for more papers by this author Christine C Jao Christine C Jao Department of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA, USA Search for more papers by this author Helen M Kent Helen M Kent MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author P Jonathan G Butler P Jonathan G Butler MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Philip R Evans Corresponding Author Philip R Evans MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Ralf Langen Corresponding Author Ralf Langen Department of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA, USA Search for more papers by this author Harvey T McMahon Corresponding Author Harvey T McMahon MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Jennifer L Gallop Jennifer L Gallop MRC Laboratory of Molecular Biology, Cambridge, UKPresent address: Department of Systems Biology, Harvard Medical School, Boston, MA 02115, USA Search for more papers by this author Christine C Jao Christine C Jao Department of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA, USA Search for more papers by this author Helen M Kent Helen M Kent MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author P Jonathan G Butler P Jonathan G Butler MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Philip R Evans Corresponding Author Philip R Evans MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Ralf Langen Corresponding Author Ralf Langen Department of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA, USA Search for more papers by this author Harvey T McMahon Corresponding Author Harvey T McMahon MRC Laboratory of Molecular Biology, Cambridge, UK Search for more papers by this author Author Information Jennifer L Gallop1,‡, Christine C Jao2,‡, Helen M Kent1, P Jonathan G Butler1, Philip R Evans 1, Ralf Langen 2 and Harvey T McMahon 1 1MRC Laboratory of Molecular Biology, Cambridge, UK 2Department of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA, USA ‡These authors contributed equally to this work *Corresponding authors: MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, UK. Tel.: +44 1223 402311; Fax: +44 1223 402310; E-mail: [email protected] of Biochemistry and Molecular Biology, Zilkha Neurogenetic Institute, University of Southern California, 1501 San Pablo Street, Los Angeles, CA 90033, USA. E-mail: [email protected] Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, UK. E-mail: [email protected] The EMBO Journal (2006)25:2898-2910https://doi.org/10.1038/sj.emboj.7601174 Present address: Department of Systems Biology, Harvard Medical School, Boston, MA 02115, USA PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Endophilin-A1 is a BAR domain-containing protein enriched at synapses and is implicated in synaptic vesicle endocytosis. It binds to dynamin and synaptojanin via a C-terminal SH3 domain. We examine the mechanism by which the BAR domain and an N-terminal amphipathic helix, which folds upon membrane binding, work as a functional unit (the N-BAR domain) to promote dimerisation and membrane curvature generation. By electron paramagnetic resonance spectroscopy, we show that this amphipathic helix is peripherally bound in the plane of the membrane, with the midpoint of insertion aligned with the phosphate level of headgroups. This places the helix in an optimal position to effect membrane curvature generation. We solved the crystal structure of rat endophilin-A1 BAR domain and examined a distinctive insert protruding from the membrane interaction face. This insert is predicted to form an additional amphipathic helix and is important for curvature generation. Its presence defines an endophilin/nadrin subclass of BAR domains. We propose that N-BAR domains function as low-affinity dimers regulating binding partner recruitment to areas of high membrane curvature. Introduction Endophilin A proteins have been implicated in membrane curvature generation in synapses during clathrin-mediated endocytosis as they bind to the endocytic proteins dynamin and synaptojanin. In Drosophila and in Caenorhabditis elegans, endophilin mutants have defective synaptic vesicle recycling (Guichet et al, 2002; Rikhy et al, 2002; Verstreken et al, 2002; Schuske et al, 2003). In higher organisms, overexpression of the endophilin SH3 domain, antibodies against endophilin and peptides that bind to the SH3 domain all result in the inhibition of vesicle recycling and the accumulation of clathrin-coated profiles, suggesting an involvement in clathrin-coated vesicle formation (Ringstad et al, 1999; Simpson et al, 1999; Gad et al, 2000). There is still some vesicle endocytosis in endophilin-deficient flies, and the slower kinetics of this residual component is consistent with clathrin-mediated endocytosis (Dickman et al, 2005). Thus, endophilin must either speed up a clathrin-mediated pathway in flies or be involved in a separate, clathrin-independent endocytic pathway that is faster than clathrin-dependent endocytosis. The ability to effect membrane curvature may implicate endophilin in early stages of vesicle formation where it could help to generate the initial membrane curvature, or in late stages where it could aid in vesicle neck formation. The stage of action has been examined in the context of clathrin-coated vesicle formation given that clathrin-coated profiles are easily observed by electron microscopy. Evidence against an early stage function for endophilin comes from studies on clathrin-coated vesicle formation in a cell-free assay. Here depletion of endophilin did not affect the number and morphology of clathrin-coated pits (Ringstad et al, 1999). A lipid-modifying activity of endophilin to aid in membrane curvature has also been excluded (Gallop et al, 2005). Evidence for late stage involvement has been obtained from this same cell-free coating assay where a significant reduction of dynamin-coated structures following endophilin depletion was observed. Thus, endophilin could be involved in late stages of endocytosis through its recruitment of dynamin and/or the lipid phosphatase synaptojanin. PtdIns(4,5)P2 is an important lipid in anchoring a number of clathrin-coated vesicle components to the membrane, including the clathrin recruitment and polymerising protein AP180 (Ford et al, 2001), and thus depletion of this lipid by the mobilisation of synaptojanin to coated vesicles would help release the coat components. Indeed, endophilin is required in C. elegans for the recruitment of synaptojanin to nerve terminals (Schuske et al, 2003), and deletion of synaptojanin in mice leads to an accumulation of coated vesicle profiles (Cremona et al, 1999). In the lamprey reticulospinal synapse, disruption of the endophilin SH3 domain interactions perturbs uncoating of clathrin-coated vesicles (Gad et al, 2000). Defective vesicle scission in this study also points to a role in dynamin recruitment. We should note however that there is no firm biochemical assignment of endophilin to clathrin-mediated endocytosis, as endophilin does not enrich in clathrin-coated vesicles nor bind to specific components of the clathrin-coat machinery, and the phenotypes observed could well be indirect. By sequence analysis, there are A and B subfamilies of endophilins. In the A subfamily, there are endophilins A1 (also called endophilin1, SH3P4, SH3GL2 and EEN-B1), A2 (also called endophilin2, SH3P8 and SH3GL1) and A3 (also called endophilin3, SH3P13 and SH3GL3) and in the B subfamily there are endophilins B1 (also called SH3GLB1) and B2 (also called Bif1, SH3GLB2 and EEN). Some of these are synaptically enriched, whereas others are more ubiquitously expressed (for review see Huttner and Schmidt, 2000). They all have the same overall domain structure, with an N-terminal N-BAR domain (BAR domain with an additional predicted N-terminal amphipathic helix) coupled to an SH3 domain by a variable linker region. The ubiquitous distribution of some endophilins, their interactions with membranes and trafficking proteins, and the role of endophilin-A1 in synaptic vesicle endocytosis support the hypothesis that endophilins perform a general function in forming transport carriers in different trafficking pathways. A homologous protein, amphiphysin, has a similar overall domain structure (with an N-BAR domain followed by an SH3 domain; see Figure 1A for scheme) and is implicated in T-tubule formation in muscle and in clathrin-coated vesicle formation (Bauerfeind et al, 1997; Razzaq et al, 2001; Lee et al, 2002; Evergren et al, 2004). Figure 1.Ordering of N-terminal residues of endophilin on membrane binding. (A) Domain structure of endophilin, nadrin and amphiphysin. The C-terminal region of nadrin has been truncated. (B) In the CD spectrum (room temperature), there is additional α-helical structure in the N-BAR domain on incubation with 50 nm Folch liposomes. This was not seen for the BAR alone (not shown). (C) EPR spectra of endophilin A1 N-BAR domains in the absence (black dash) and presence (red) of liposomes. Sample traces for residues 2, 4, 5 and 10 are shown. Other traces are shown in Supplementary Figure 1. Asterisks point to additional immobilisation compared to surrounding residues on membrane binding. Protein (2 μM) was incubated with 1.4 mg/ml liposomes and centrifuged to separate bound from unbound. Membrane-bound spectra are shown at a magnification of 2.5. Download figure Download PowerPoint The deformation of membrane that is required to make small-diameter transport vesicles, as found at synapses, has a significant energetic requirement. When making small liposomes in vitro, this energy is provided by intense sonication. In vivo, high curvature can be achieved using peripheral membrane binding proteins that effect and stabilise curvature (for review see McMahon and Gallop, 2005). In particular, the insertion of amphipathic helices into the hydrophobic phase of the bilayer is proposed to be a general biophysical mechanism for curvature generation during vesicle budding, based on point mutagenesis in amphiphysin (Peter et al, 2004), endophilin (Farsad et al, 2001), epsin (Ford et al, 2002) and Sar1 GTPase (Lee et al, 2005). Until now, insertion of amphipathic helices for vesicle budding proteins has not been shown directly. Here we show, using electron paramagnetic resonance spectroscopy (EPR), that the N-terminal amphipathic helix of endophilin inserts into membranes and we elucidate the orientation and depth of helix penetration. In the case of N-BAR domains, both their amphipathic helices and BAR domains have been implicated in the promotion of membrane curvature, and the relative importance of these two modules has been unclear. In vitro, the N-BAR domain of endophilin tubulates liposomes and a truncation that includes approximately half the BAR domain is also effective, as are isolated BAR domains (Farsad et al, 2001; Peter et al, 2004). We now carry out a thorough analysis of the endophilin N-BAR domain using crystallographic and biophysical techniques. The principles uncovered (driving of curvature by amphipathic helices and selection or limiting of membrane curvature by a BAR domain, and the feed-forward behaviour of N-BAR domain binding) will also apply to other proteins where one finds this same combination of amphipathic helix followed by BAR domain, including nadrin/RICH and BRAP1/Bin2 N-BAR domains. By homology screening, we also find that nadrin and endophilin are in the same structural subclass of BAR domain owing to an insert present on the concave face. This insert includes a further predicted amphipathic helix that exhibits membrane interaction capability. Our measurements of the behaviour of amphipathic helices on membranes are also likely true for the epsin family of proteins and also for the Arf, Arl and Sar family GTPases. Results An N-terminal amphipathic helix of endophilin folds and inserts into membranes Predicted N-terminal amphipathic helices have been proposed to fold on membrane binding and anchor membrane curvature-generating proteins in the membrane to cause displacement of lipids in one leaflet, promoting curvature generation (Ford et al, 2002; Peter et al, 2004). We now use direct biophysical methods to determine the structure of these residues on membrane binding. On liposome binding, there is an increase in α-helicity of the N-BAR domain of endophilin from 36 to 48% (estimated from circular dichroism; see Figure 1B), which is not observed in the absence of the N-terminal residues (not shown), implying the formation of additional helical structures. To test how the predicted N-terminal amphipathic helix folds, whether it inserts into membranes and to determine its topology in relation to the membrane, we used EPR together with site-directed spin labelling. A series of N-BAR domains were made where cysteines were substituted for each residue from 2 to 16 by site-directed mutagenesis. Spin labels were then attached to each cysteine mutant and the protein was used for EPR analysis. The EPR spectra of the spin-labelled derivatives in solution are very sharp, and on membrane binding these are broadened for each residue. These data show that this N-terminal region is disordered in solution and becomes ordered upon membrane binding (see example spectra in Figure 1C and the full range in Supplementary Figure 1). Residues 4, 5, 8 and 16, which are predicted to lie on a single plane of the predicted amphipathic helix, also show additional immobilisation (see asterisks in Figure 1C). This may indicate some other interactions with the BAR domain or lipid headgroups. The accessibility of each spin-labelled site to oxygen (which preferentially partitions into membranes) and to NiEDDA (preferentially in solution) is plotted in Figure 2A and shows which residues penetrate the bilayer. Up to residue 16, the O2 and NiEDDA accessibilities (Π(O2) and Π(NiEDDA)) exhibit a periodicity of 3–4 amino acids, consistent with the formation of a continuous α-helical structure. Importantly, the periodicities of access to the respective colliders are 180° out of phase. Such behaviour is typical for asymmetrically solvated α-helices in which one face is exposed to the membrane where the accessibility to O2 is high; in contrast, residues on the opposing face are solvent-exposed and consequently more accessible to the hydrophilic NiEDDA. The membrane exposure of a given site can be conveniently summarised by the contrast parameter Φ=ln(Π(O2)/Π(NiEDDA)), which is proportional to the depth of membrane immersion (Altenbach et al, 1994). As shown in Figure 2B and C, membrane-exposed residues, corresponding to local maxima of Φ, cluster on one side of the helical wheel, whereas solvent-exposed sites (local minima of Φ) lie on the opposite face. The polar, more solvent-exposed residues of the helix have considerable positive charge and will prefer negatively charged lipid headgroups or a negative patch on an adjacent protein. The immersion depths of the lipid-facing residues were calibrated using labelled hydrocarbon chains (Altenbach et al, 1994) (see Materials and methods). Based on this calibration and the data in Figure 2A, we can place the centre of the helix near the phosphate level (Figure 2D). This is the first direct demonstration of amphipathic helix membrane insertion for an endocytic protein and we propose that this will apply to all N-BAR proteins, epsin family members and Arf/Arl/Sar proteins, thus providing a potential general mechanism by which membrane curvature is generated by these classes of proteins. Figure 2.Membrane insertion and orientation of endophilin N-terminal amphipathic helix. (A) Oxygen (red circles) and NiEDDA (green squares) accessibilities (Π) of membrane-bound N-BAR domain as a function of label position. The graph below shows a ln(Π ratio) plot (Φ) showing the differential access of colliders to the spin label and the penetration of hydrophobic residues into the membrane. The periodic oscillation is indicative of a helical structure. Equivalent maxima indicate that the helix lies planar to the membrane. (B) Helical wheel representation showing hydrophobic and charged faces. (C) Model of the amphipathic helix, residues 1–16 with hydrophobic residues coloured green and surface charge potential also shown. (D) Model of the N-BAR amphipathic helix to scale with PtdIns(4,5)P2 and PtdSer lipids showing the depth of penetration of the helix as calculated from data in (A) and penetration measurements, described in Materials and methods. Download figure Download PowerPoint BAR domain structure The N-terminal amphipathic helix of endophilin is followed immediately by a predicted BAR domain, which is expected to sense or stabilise positive membrane curvature (Peter et al, 2004). As the sequence homology between the endophilin and amphiphysin BAR domains (where a structure was already available) is low, we crystallised rat endophilin BAR to elucidate the curvature of the domain (Figure 3A and Supplementary Figure 2). A similar structure of mouse endophilin BAR domain has since been published (Weissenhorn, 2005) and when these are overlaid the r.m.s. deviation on the dimer is 0.95 Å for 392/408 residues. Structural details are marked on the endophilin sequence in Figure 4 and the N-terminal amphipathic helix is labelled as helix zero (after the epsin ENTH nomenclature; Ford et al, 2002) given that it folds on membrane binding (Figure 2A) despite being invisible in the unliganded crystal structure. The surface charge distribution of endophilin is similar to that of amphiphysin but more negative charges are concentrated on the convex face (see endophilin in Figure 3C and amphiphysin in Figure 3D). The high negative charge on the convex surface is conserved among endophilins (Supplementary Figure 2). In the structure solved by Weissenhorn, 11 cadmium ions were bound to the surface and he posits that these may mimic calcium binding sites of endophilin in vivo. We looked for calcium binding using di-bromo BAPTA, which allows detection of low micromolar affinities, and by using isothermal titration calorimetry, which allows detection of nanomolar affinity interactions (Figure 3F and G). We see no evidence of specific calcium interactions. Figure 3.Endophilin N-BAR crystal structure. (A) Ribbon diagram of the banana-shaped rat endophilin-A1 BAR domain (Protein Data Bank (PDB) accession number 2c08) with a view of the concave surface below. Monomers are dark to light from NH2- to COOH-termini with one coloured in brown to yellow and the other in dark blue to light blue. Lysine and arginine residues potentially important for membrane binding and K227 used to examine dimerisation are marked. (B) Superposition of the endophilin BAR domain (orange) with amphiphysin BAR (green). (C) Surface representation of the BAR domain of endophilin coloured according to electrostatic potential and a mesh equipotential surface contoured at 0.05 V. (D) Similar representation for amphiphysin as in (C). Both molecules are negatively charged (red) except on the concave face and the tips of the crescent, which are positively charged (blue). The overall shapes are very similar. (E) Hydrophobic residues in the dimer interface of rat endophilin-A1 BAR are coloured green on the surface-represented monomer. Dimensions of the BAR domain are also indicated. (F) Calcium does not bind to the endophilin N-BAR domain. The endophilin N-BAR domain was decalcified by purification in the presence of 2 mM EDTA followed by incubation with 10 mM EDTA and then extensive dialysis against Ca-free buffer (prepared using plastics). The absorbance of di-bromo BAPTA (affinity for Ca2+ of 2 μM in the absence of Mg2+) at 265 nm was followed on calcium titration in the presence and absence of decalcified endophilin BAR domain. No difference was observed, suggesting that endophilin does not effectively compete with di-bromo BAPTA, even at double the concentration, for Ca2+ ions. (G) Isothermal titration calorimetry was used to test for heat changes on CaCl2 injection into decalcified endophilin N-BAR domain. These experiments were performed with 6 μl injections of 1 mM CaCl2 into 45 μM full-length endophilin and 76 μM endophilin N-BAR domain at 10 or 25°C. Example results for the N-BAR domain are shown and no binding was observed. (H) Modelling of helix1 insert residues 62–79 as an α-helix. Hydrophobic residues are predominantly on one side and are flanked by positively charged residues. The remaining residues on this insert are not predicted to form an amphipathic helix. (I) EPR spectra of N-BAR M70C in the absence (black dash) and presence (red) of liposomes. The red spectrum is magnified by 2.5. Download figure Download PowerPoint Figure 4.Structure-based sequence alignment of endophilin and amphiphysin BAR domains and alignment to nadrin showing close homology to endophilin. Download figure Download PowerPoint The curvature of endophilin BAR, formed by the angle of dimerisation and kinks in its helices, is very close to that of amphiphysin BAR (see overlay in Figure 3B and structural alignment in Figure 4) and arfaptin BAR (not shown), and thus these proteins cannot be distinguished by a difference in their predicted membrane curvature preference. A large hydrophobic patch is buried in the dimer interface (coloured green on the surface-represented monomer in Figure 3E). The buried surface area is 2870 Å2 per monomer, whereas in amphiphysin only 2400 Å2 is buried. The curvature of the domain is likely to be rigid, as the mouse and the rat endophilin BAR structures are derived from crystals with completely different crystal packing, yet these structures superimpose closely and show the same radius of curvature (85 Å in the absence of the helix1 insert; Figure 3E; see next paragraph). In the present structure of the rat endophilin BAR, the extremities are involved in crystal contacts, leading to ordering of these flexible regions. Endophilin BAR has an extra insert in the middle of its membrane binding face as compared to amphiphysin BAR. We call this the ‘helix1 insert’ (H1I, residues 60–87). The sequence of this insert differs considerably between endophilins and is also found in nadrin N-BAR family members. The QPNP sequence, which follows the break in helix1, appears to be diagnostic for endophilin family members across different species (Supplementary Figure 2). The H1I is mostly invisible (and thus disordered) in the crystal structure (Figure 2A) apart from an initial short helix (see Figure 4). This is predicted to continue as an amphipathic helix for several more turns. This would be particularly favoured in the low dielectric constant environment under the BAR domain and near the membrane. In a helical wheel representation (see Figure 3H), the hydrophobic side of the predicted helix is flanked on both sides by positively charged residues. This may indicate penetration of the membrane by the hydrophobic residues (similar to the N-terminal amphipathic helix) with accompanying electrostatic interactions with the charged lipid headgroups. Although by circular dichroism we were unable to detect an increase in helicity upon addition of liposomes to the BAR domain, this is not surprising as the BAR domain alone does not bind well to membranes and at least part of the helix appears to be already folded before binding. We chose one residue beyond the initial helix (M70) to test the possibility of folding on membrane interaction. The EPR spectrum shows some ordering of this residue upon membrane interaction (Figure 3I). It should be noted, however, that the spectral change upon membrane interaction is not as pronounced as in the N-terminal regions. This is owing to the fact that, in solution, position 70 is less dynamic than the N-terminal residues and consequently the observed mobility changes are less dramatic. This is also not surprising given that this helix appears to start to fold in the crystal structure. The O2 and NiEDDA accessibilities for residue 70 resulted in a Φ value of 1.5, demonstrating that this position is indeed membrane-exposed at an immersion depth of approximately 6 Å. A predicted amphipathic sequence is also found at the C-terminal end of the nadrin insert. We find no evidence in our structure for a lysophosphatidic acid acyl transferase active site. We have previously tested extensively for biochemical evidence of this activity and showed that it was a contamination of protein preparation (Gallop et al, 2005). To test if the dimerisation seen in the crystal holds true in solution, we used equilibrium ultracentrifugation of the N-BAR domain (Figure 5A and residuals plotted in Figure 5B) and full-length protein (Supplementary Figure 4A). The dimerisation constant for the N-BAR is 10 μM and fits very well to a monomer:dimer equilibrium. This means that the protein could well be monomeric in cells—the concentration of endophilin in brain extract was estimated by blotting to be ∼0.1 μM, making the concentration at the synapse at perhaps ∼1 μM. For the full-length protein, there is evidence of higher order oligomers at high concentrations (Supplementary Figure 4A). This could be due to the previously proposed intramolecular interaction between the SH3 domain and the central proline-rich linker region (Chen et al, 2003). Figure 5.Endophilin N-BAR dimerisation on membrane binding. (A) Equilibrium sedimentation data for rat endophilin-A1 N-BAR domain. Endophilin N-BAR domain dimerises with a Kd of ∼10 μM. The readings from three cells are fitted with an ideal monomer/dimer equilibrium that fits with a 26 kDa monomer. The calculated molecular mass is 28.7 kDa. Residuals are plotted in (B). (C) EPR spectra of membrane-bound endophilin reveals dimer interactions. Spectra were obtained either from protein that was fully labelled at position 227 (red trace) or from a mixture of 25% labelled protein and 75% unlabelled protein (black trace). The scan width was 200 Gauss. The amplitudes and line shapes of the respective spectra were very different owing to the strong dipolar spin–spin interactions that were present in the fully labelled case. The difference between the respective spectra was used to determine the distances using a Pake pattern analysis (Altenbach et al, 2001), and the blue spectrum can be used to evaluate the quality of this distance analysis. It is based on simulations in which the calculated distance distribution (shown in panel D) was used to generate a dipolar broadened spectrum from the black spectrum. This simulated spectrum closely corresponds to the observed spectrum for the fully labelled case indicating a good quality fit. Download figure Download PowerPoint As the model for curvature sensing by BAR domains is binding via the concave face (Peter et al, 2004) and the concave nature is only found in the dimeric form, it is surprising that the KD for dimerisation in solution is as high as 10 μM (Figure 5A). Spin coupling between labelled site-directed cysteine mutants at position 227 was used to test if the dimer is the predominant form of the protein on membranes. K227 is located near the dimerisation interface, and the α-carbon distance between the two K227 residues in the crystal dimer is 8 Å (see residue marked in Figure 3A). Introduction of a spin label at position 227 gave rise to a strong spin–spin interaction for this endophilin mutant and the resulting spectrum of the membrane-bound form exhibited strong dipolar broadening that is characteristic for spin labels in close proximity (Figure 5C, red trace). This spectrum was very different from a control spectrum for the K227 derivative, in which the dipolar interaction was strongly reduced by co-mixing of 25% labelled protein with 75% unlabelled protein (Figure 5C, black trace). Quantitative analysis of the spectra with and without dipolar interaction allowed us to determine inter-spin label distances using Pake patterns (Rabenstein and Shin, 1995; Altenbach et al, 2001) and the resulting distances ranged from ∼8 to 10 Å (Figure 5D). These data are in excellent agreement with the distance between these labels predicted from the crystal structure and clearly demonstrate that membrane-bound endophilin forms the dimer interactions seen in the crystal. Amphipathic helices and the BAR domain of endophilin collaborate to effect membrane curvature The ability of endophilin to generate and stabilise membrane curvature can be assessed using liposomes rich in negatively charged lipids to which the N-BAR domain binds. By electron microscopy, we can determine the shape changes of these liposomes. The amphiphysin N-BAR domain and the arfaptin2 BAR domain constrict liposomes into tubules and higher concentrations of the BAR dom
