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Fusogenic Tilted Peptides Induce Nanoscale Holes in Supported Phosphatidylcholine Bilayers Karim El Kirat,† Laurence Lins,‡ Robert Brasseur,‡ and Yves F. Dufreˆne*,† Unite´ de chimie des interfaces, Universite´ catholique de Louvain, Croix du Sud 2/18, B-1348 Louvain-la-Neuve, Belgium, and Centre de Biophysique Mole´ culaire Nume´ rique, Faculte´ Universitaire des Sciences Agronomiques de Gembloux, Passage des De´ porte´ s, 2, B-5030 Gembloux, Belgium Received September 22, 2004. In Final Form: January 13, 2005 Tilted peptides are known to insert in lipid bilayers with an oblique orientation, thereby destabilizing membranes and facilitating membrane fusion processes. Here, we report the first direct visualization of the interaction of tilted peptides with lipid membranes using in situ atomic force microscopy (AFM) imaging. Phase-separated supported dioleoylphosphatidylcholine/dipalmitoylphosphatidylcholine (DOPC/DPPC) bilayers were prepared by fusion of small unilamellar vesicles and imaged in buffer solution, in the absence and in the presence of the simian immunodeficiency virus (SIV) peptide. The SIV peptide was shown to induce the rapid appearance of nanometer scale bilayer holes within the DPPC gel domains, while keeping the domain shape unaltered. We attribute this behavior to a local weakening and destabilization of the DPPC domains due to the oblique insertion of the peptide molecules. These results were directly correlated with the fusogenic activity of the peptide as determined using fluorescently labeled DOPC/DPPC liposomes. By contrast, the nontilted ApoE peptide did not promote liposome fusion and did not induce bilayer holes but caused slight erosion of the DPPC domains. In conclusion, this work provides the first direct evidence for the production of stable, well-defined nanoholes in lipid bilayer domains by the SIV peptide, a behavior that we have shown to be specifically related to the tilted character of the peptide. A molecular mechanism underlying spontaneous insertion of the SIV peptide within lipid bilayers and the subsequent removal of bilayer patches is proposed, and its relevance to membrane fusion processes is discussed.
Introduction Membrane fusion is a key process for cell life and development.1-3 Many fusion events are known to involve the active participation of hydrophobic peptides, which help destabilize the membrane lipid bilayer.4 Tilted peptides5 represent a special class of fusogenic peptides found in many membrane-interacting proteins such as viral fusion proteins, neurotoxic proteins, and proteins involved in lipoprotein metabolism.6 These short peptides (10-20 residues) have a hydrophobicity gradient that runs along the axis of their helical structure. Hence, not only are they amphipathic but their hydrophobicity increases from one end of the helix to the other, a property that causes them to insert at an angle of 30-60° at hydrophobic/ hydrophilic interfaces.7 Both theoretical and experimental methods have been used to investigate tilted peptides. Using molecular modeling, the envelope protein of the Newcastle disease virus (NDV) was the first tilted peptide to be discovered.8 Since then, many different tilted peptides have been detected in other proteins, including the fusion protein of * Corresponding author. Phone: (32) 10 47 36 00. Fax: (32) 10 47 20 05. E-mail:
[email protected]. † Universite ´ catholique de Louvain. ‡ Faculte ´ Universitaire des Sciences Agronomiques de Gembloux. (1) Peuvot, J.; Schanck, A.; Lins, L.; Brasseur, R. J. Theor. Biol. 1999, 198, 173-181. (2) Jena, B. P. J. Cell. Mol. Med. 2004, 8, 1-21. (3) Malinin, V. S.; Lentz, B. R. Biophys. J. 2004, 86, 2951-2964. (4) Chizmadzhev, Y. A. Bioelectrochemistry 2004, 63, 129-136. (5) Brasseur, R. Mol. Membr. Biol. 2000, 17, 31-40. (6) Brasseur, R.; Pillot, T.; Lins, L.; Vandekerckhove, J.; Rosseneu, M. Trends Biochem. Sci. 1997, 22, 167-171. (7) Lins, L.; Flore, C.; Chapelle, L.; Talmud, P. J.; Thomas, A.; Brasseur, R. Protein Eng. 2002, 15, 513-520. (8) Brasseur, R.; Lorge, P.; Goormaghtigh, E.; Ruysschaert, J. M.; Espion, D.; Burny, A. Virus Genes 1988, 1, 325-332.
the Simian immunodeficiency virus (SIV), the prion protein, the β-amyloid peptide, lipases, and some lipoproteins.6 Molecular modeling predicted an oblique insertion of these peptides in lipid bilayers, while experimental assays using liposomes have demonstrated that they induce the fusion of lipid membranes.7-9 Furthermore, Fourier transform infrared (FTIR) spectroscopy has allowed theoretical predictions concerning the average oblique insertion of tilted peptides of viral fusion proteins in lipid bilayers to be confirmed.10 Finally, the existence of tilted peptides in lipid bilayers has also been confirmed by neutron diffraction experiments.11 Despite the vast body of literature that has accumulated on tilted peptides in the past decade, in situ high resolution visualization of their interaction with lipid membranes has never been reported. Atomic force microscopy (AFM) is a powerful tool for imaging biological specimens at high resolution.12-14 A key advantage of the technique over electron microscopy is that it can work in aqueous solution, thereby making it possible to probe the sample in its native state. In recent years, rapid advances have been made in applying AFM to supported lipid membranes, indicating that the technique is taking root in the lipid film research com(9) Pillot, T.; Lins, L.; Goethals, M.; Vanloo, B.; Baert, J.; Vandekerckhove, J.; Rosseneu, M.; Brasseur, R. J. Mol. Biol. 1998, 274, 381393. (10) Martin, I.; Dubois, M. C.; Defrise-Quertain, F.; Saermark, T.; Burny, A.; Brasseur, R.; Ruysschaert, J. M. J. Virol. 1994, 68, 11391148. (11) Bradshaw, J. P.; Darkes, M. J.; Harroun, T. A.; Katsaras, J.; Epand, R. M. Biochemistry 2000, 39, 6581-6585. (12) Morris, V. J.; Kirby, A. R.; Gunning, A. P. Atomic Force Microscopy for Biologists; Imperial College Press: London, 1999. (13) Engel, A.; Muller, D. J. Nat. Struct. Biol. 2000, 7, 715-718. (14) Jena, B. P.; Ho¨rber, J. K. H. Atomic Force Microscopy in Cell Biology, Methods in Cell Biology; Academic Press: San Diego, CA, 2002; Vol. 68.
10.1021/la047640q CCC: $30.25 © 2005 American Chemical Society Published on Web 02/23/2005
AFM of Tilted Peptide-Membrane Interactions
munity.15,16 AFM has been used to observe, in aqueous solution, the effect of various external agents such as solvents, peptides, and antibiotics on the structure of supported lipid films. In early work, Mou and co-workers17 showed that low percentages of ethanol induce interdigitation in dipalmitoylphosphatidylcholine (DPPC) supported bilayers. The same author also showed that the interaction between gramicidin A and DPPC bilayers leads to the formation of highly ordered domains.18 Rinia and co-workers provided evidence for the formation of striated domains of lytic peptides within DPPC bilayers.19,20 Filipininduced lesions were visualized in lipid membranes containing cholesterol.21,22 Yip and McLaurin23 observed the formation of fibrils by the β-amyloid peptide in association with lipid bilayers. The amphipathic membrane-active peptide melittin was shown to cause dramatic disruption of lipid bilayers.24 Ha and co-workers25 have described the disruption of a dipalmitoylphosphatidic acid bilayer induced by the antimicrobial tridecapeptide indolicidin. In this paper, we report the direct observation of the interaction of tilted peptides with supported lipid membranes using in situ AFM imaging. We chose the SIV fusion peptide because it is one of the best characterized tilted peptides,7,10,11,26-33 has a high fusogenic activity, and is thought to destabilize lipid membranes by disturbing the parallel organization of the lipid acyl chains.34 As a control, we used the amphipathic ApoE peptide which is responsible for the interaction of the whole apolipoprotein E with the lipoprotein surface without any fusogenic activity. Mixed dioleoylphosphatidylcholine/dipalmitoylphosphatidylcholine (DOPC/DPPC) bilayers were used as model membranes because choline phospholipids are commonly found in eukaryotic cell membranes and this specific mixture is known to separate at room temperature into (15) Dufreˆne, Y. F.; Lee, G. U. Biochim. Biophys. Acta 2000, 1509, 14-41. (16) Janshoff, A.; Steinem, C. ChemBioChem 2001, 2, 798-808. (17) Mou, J.; Yang, J.; Huang, C.; Shao, Z. Biochemistry 1994, 33, 9981-9985. (18) Mou, J.; Czajkowsky, D. M.; Shao, Z. Biochemistry 1996, 35, 3222-3226. (19) Rinia, H. A.; Kik, R. A.; Demel, R. A.; Snel, M. M.; Killian, J. A.; van Der Eerden, J. P.; de Kruijff, B. Biochemistry 2000, 39, 5852-5858. (20) Rinia, H. A.; Boots, J. W.; Rijkers, D. T.; Kik, R. A.; Snel, M. M.; Demel, R. A.; Killian, J. A.; van der Eerden, J. P.; de Kruijff, B. Biochemistry 2002, 41, 2814-2824. (21) Santos, N. C.; Ter-Ovanesyan, E.; Zasadzinski, J. A.; Prieto, M.; Castanho, M. A. Biophys. J. 1998, 75, 1869-1873. (22) Lawrence, J. C.; Saslowsky, D. E.; Edwardson, J. M.; Henderson, R. M. Biophys. J. 2003, 84, 1827-1832. (23) Yip, C. M.; McLaurin, J. Biophys. J. 2001, 80, 1359-1371. (24) Steinem, C.; Galla, H. J.; Janshoff, A. Phys. Chem. Chem. Phys. 2000, 2, 4580-4585. (25) Ha, T. H.; Kim, C. H.; Park, J. S.; Kim, K. Langmuir 2000, 16, 871-875. (26) Martin, I.; Defrise-Quertain, F.; Mandieau, V.; Nielsen, N. M.; Saermark, T.; Burny, A.; Brasseur, R.; Ruysschaert, J. M.; Vandenbranden, M. Biochem. Biophys. Res. Commun. 1991, 175, 872-879. (27) Horth, M.; Lambrecht, B.; Khim, M. C.; Bex, F.; Thiriart, C.; Ruysschaert, J. M.; Burny, A.; Brasseur, R. EMBO J. 1991, 10, 27472755. (28) Voneche, V.; Portetelle, D.; Kettmann, R.; Willems, L.; Limbach, K.; Paoletti, E.; Ruysschaert, J. M.; Burny, A.; Brasseur, R. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 3810-3814. (29) Schanck, A.; Peuvot, J.; Brasseur, R. Biochem. Biophys. Res. Commun. 1998, 250, 12-14. (30) Martin, I.; Dubois, M. C.; Saermark, T.; Epand, R. M.; Ruysschaert, J. M. FEBS Lett. 1993, 333, 325-330. (31) Epand, R. F.; Martin, I.; Ruysschaert, J. M.; Epand, R. M. Biochem. Biophys. Res. Commun. 1994, 205, 1938-1943. (32) Colotto, A.; Martin, I.; Ruysschaert, J. M.; Sen, A.; Hui, S. W.; Epand, R. M. Biochemistry 1996, 35, 980-989. (33) Martin, I.; Turco, S. J.; Epand, R. M.; Ruysschaert, J. M. Eur. J. Biochem. 1998, 258, 150-156. (34) Lins, L.; Charloteaux, B.; Thomas, A.; Brasseur, R. Proteins 2001, 44, 435-447.
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fluid and gel phases.35 The influence of the solvent trifluoroethanol, usually used to increase the peptide solubility in aqueous solution, was investigated, and the ability of the peptides to perturb the nanoscale organization of the supported bilayers was correlated with their fusogenic activity using fluorescently labeled liposomes. The results point to the power of AFM for elucidating peptidemembrane interactions at high resolution and, in turn, the molecular basis of fusion processes. Experimental Section Materials. L-R-Dioleoylphosphatidylcholine (DOPC) and L-Rdipalmitoylphosphatidylcholine (DPPC) were purchased from Sigma (St. Louis, MO). Octadecyl rhodamine chloride (R18) was obtained from Molecular Probes (Eugene, OR). SIV (NH2GVFVLGFLGFLA-CONH2) and ApoE (Ac-EDMQRQWAGLVEKVQAA-CONH2) peptides were obtained from Neosystem (Strasbourg, France). They were 90% pure. Other chemicals were purchased from Merck (Darmstadt, Germany). Stock solutions of peptides were prepared in trifluoroethanol (TFE) at a 600 µM final concentration. For AFM experiments, stock solutions were diluted into Tris/EDTA buffer (10 mM Tris, 150 mM NaCl, and 5 mM EDTA; pH 7.4) to a final peptide concentration of 10 µM. For assays in the absence of TFE, the organic solutions were dried under nitrogen and then the peptides were resuspended in TBS/EDTA buffer by vigorous agitation and smooth sonication (5 min in a 45 kHz sonicating bath). Preparation of Supported Lipid Bilayers. Supported lipid bilayers were prepared using the vesicle fusion method.35,36 To this end, DOPC and DPPC were dissolved in chloroform at a 1 mM final concentration. An equimolar mixture of these two lipids was then evaporated under nitrogen and dried in a desiccator under vacuum for 2 h. Multilamellar vesicles (MLVs) were obtained by resuspending the lipidic dried film in calciumcontaining buffer (10 mM Tris, 150 mM NaCl, and 3 mM CaCl2; pH 7.4) at a 1 mM final lipid concentration. To obtain small unilamellar vesicles (SUVs), the suspension was sonicated to clarity (four cycles of 2 min) using a 500 W probe sonicator (Fisher Bioblock Scientific, France; 35% of the maximal power; 13 mm probe diameter) while keeping the suspension in an ice bath. The liposomal suspension was then filtered on 0.2 µm nylon filters (Whatman Inc., U.S.A.) to eliminate titanium particles. Freshly cleaved mica squares (16 mm2) were glued onto steel sample pucks (Veeco Metrology LLC, Santa Barbara, CA) using Epotek 377 (Gentec Benelux, Waterloo, Belgium). A 2 mL portion of the SUV suspension was then deposited onto the mica samples, and the SUVs were allowed to adsorb and fuse on the solid surface for 1 h at 60 °C. Subsequently, the sample was rinsed five times with 2 mL of EDTA-containing buffer (10 mM Tris, 150 mM NaCl, and 5 mM EDTA; pH 7.4) to remove Ca2+ ions. The samples were then slowly cooled to room temperature. Atomic Force Microscopy. Supported bilayers were investigated using a commerical atomic force microscope (NanoScope IV MultiMode AFM, Veeco Metrology LLC, Santa Barbara, CA) equipped with a 125 µm × 125 µm × 5 µm scanner (J-scanner). AFM images were obtained in contact mode at room temperature (23-25 °C) either in Tris/EDTA buffer (10 mM Tris, 150 mM NaCl, and 5 mM EDTA; pH 7.4) or after ∼10 min of incubation in Tris/EDTA buffer containing 10 µM peptide. All images were recorded using oxide-sharpened microfabricated Si3N4 cantilevers (Microlevers, Veeco Metrology LLC, Santa Barbara, CA) with a spring constant of 0.01 N/m (manufacturer specified), with a minimal applied force (
