Local Membrane Mechanics of Pore-Spanning Bilayers Ingo Mey, † Milena Stephan, † Eva K. Schmitt, ‡ Martin Michael Mu ¨ ller, § Martine Ben Amar, § Claudia Steinem, ‡ and Andreas Janshoff* ,† Institute of Physical Chemistry, UniVersity of Go ¨ttingen, Tammannstrasse 6, 37077 Go ¨ttingen, Germany, Institute of Organic and Biomolecular Chemistry, UniVersity of Go ¨ttingen, Tammannstrasse 2, 37077 Go ¨ttingen, Germany, and Laboratoire de Physique Statistique de l’Ecole Normale Supe ´rieure (UMR 8550), associe ´ aux UniVersite ´s Paris 6 et Paris 7 et au CNRS; 24, rue Lhomond, 75005 Paris, France Received November 24, 2008; E-mail: ajansho@gwdg.de Abstract: The mechanical behavior of lipid bilayers spanning the pores of highly ordered porous silicon substrates was scrutinized by local indentation experiments as a function of surface functionalization, lipid composition, solvent content, indentation velocity, and pore radius. Solvent-containing nano black lipid membranes (nano-BLMs) as well as solvent-free pore-spanning bilayers were imaged by fluorescence and atomic force microscopy prior to force curve acquisition, which allows distinguishing between membrane- covered and uncovered pores. Force indentation curves on pore-spanning bilayers attached to functionalized hydrophobic porous silicon substrates reveal a predominately linear response that is mainly attributed to prestress in the membranes. This is in agreement with the observation that indentation leads to membrane lysis well below 5% area dilatation. However, membrane bending and lateral tension dominate over prestress and stretching if solvent-free supported membranes obtained from spreading giant liposomes on hydrophilic porous silicon are indented. An elastic regime diagram is presented that readily allows determining the dominant contribution to the mechanical response upon indentation as a function of load and pore radius. Introduction Membrane mechanics plays a pivotal role in many biological processes such as exo- and endocytosis and cell shape/volume regulations. The ability of a lipid bilayer to deform or dilate strongly depends on its chemical composition. To assess quantitative data on the elastic properties of lipid bilayers, mainly nonlocal methods such as micropipet aspiration of giant liposomes have been used. 1,2 However, due to the mosaic nature of biological membranes, it is highly desirable to measure local elastic properties of lipid bilayers on submicrometer length scales. So far, the lack of suitable model systems prevented the use of scanning force techniques to map the elastic properties of membranes in a defined manner. Conventional solid supported lipid bilayers can only be compressed, 3 but bending or stretching requires a second aqueous compartment. With the advent of pore-spanning lipid bilayers, new stable membrane models became available that allow addressing nano- to micrometer sized free-standing membrane patches organized in a well- defined array. 4-9 While classical black lipid membranes (BLMs) lack stability and addressability with local probe microscopy, nano-BLMs - a hybrid between solid supported membranes and BLMs - offer unprecedented mechanical stability over days with mesh sizes between 20 nm and several micrometers in a defined geometric pattern. 10-12 In previous publications, it has been shown that force indentation curves can easily be acquired from the center of the pore. For electrostatically adsorbed N,N- dioctadecyl-N,N-dimethylammonium bromide (DODAB) bilay- ers on mercaptopropionic acid-coated gold-covered porous alumina, we found a rather linear behavior of the force indentation curve, which is dominated by lateral tension, 13 whereas Scheuring and co-workers 14 observed a nonlinear force response attributed to stretching at higher load. Lorenz et al. 15 † Institute of Physical Chemistry, University of Go ¨ttingen. ‡ Institute of Organic and Biomolecular Chemistry, University of Go ¨ttingen. § Laboratoire de Physique Statistique de l’Ecole Normale Supe ´rieure (UMR 8550). (1) Evans, E.; Heinrich, V.; Ludwig, F.; Rawicz, W. Biophys. J. 2003, 85, 2342–2350. (2) Rawicz, W.; Olbrich, K. C.; McIntosh, T.; Needham, D.; Evans, E. Biophys. J. 2000, 79, 328–339. (3) Künneke, S.; Krüger, D.; Janshoff, A. Biophys. J. 2004, 86, 1545– 1553. (4) Hennesthal, C.; Drexler, J.; Steinem, C. ChemPhysChem 2002, 3, 885– 889. (5) Hennesthal, C.; Steinem, C. J. Am. Chem. Soc. 2000, 122, 8085–8086. (6) Drexler, J.; Steinem, C. J. Phys. Chem. B 2003, 107, 11245–11254. (7) Favero, G.; Capanella, L.; Cavallo, S.; D’Annibale, A.; Perrella, M.; Mattei, E.; Ferri, T. J. Am. Chem. Soc. 2005, 127, 8103-8111. (8) Hemmler, R.; Bose, G.; Wagner, R.; Peters, R. Biophys. J. 2005, 88, 4000–4007. (9) Han, X. J.; Studer, A.; Sehr, H.; Geissbuhler, I.; Di Berardino, M.; Winkler, F. K.; Tiefenauer, L. X. AdV. Mater. 2007, 19, 4466–4470. (10) Ro ¨mer, W.; Lam, Y. H.; Fischer, D.; Watts, A.; Fischer, W. B.; Go ¨ring, P.; Wehrspohn, R. B.; Go ¨sele, U.; Steinem, C. J. Am. Chem. Soc. 2004, 126, 16267–16274. (11) Ro ¨mer, W.; Steinem, C. Biophys. J. 2004, 86, 955–965. (12) Schmitt, E. K.; Vrouenraets, M.; Steinem, C. Biophys. J. 2006, 91, 2163–2171. (13) Steltenkamp, S.; Müller, M. M.; Deserno, M.; Hennesthal, C.; Steinem, C.; Janshoff, A. Biophys. J. 2006, 91, 217–226. (14) Goncalves, R. P.; Agnus, G.; Sens, P.; Houssin, C.; Bartenlian, B.; Scheuring, S. Nat. Methods 2006, 3, 1007–1012. (15) Lorenz, B. M.; Mey, I.; Steltenkamp, S.; Fine, T.; Rommel, C.; Mu ¨ller, M. M.; Maiwald, A.; Wegener, J.; Steinem, C.; Janshoff, A. Small 2009, 5, 832-838. Published on Web 05/04/2009 10.1021/ja809165h CCC: $40.75  2009 American Chemical Society J. AM. CHEM. SOC. 2009, 131, 7031–7039 9 7031