Advertisement

Probing Single Membrane Proteins by Atomic Force Microscopy

  • S. Scheuring
  • K. Tanuj Sapra
  • Daniel J. Müller
Chapter

Abstract

In this book chapter, we describe the working principle of the atomic force microscope (AFM), followed by the applications of AFM in high-resolution imaging and single-molecule force spectroscopy of membrane proteins. In the imaging mode, AFM allows observing the assembly of membrane proteins directly in native membranes approaching a resolution of ~0.5 nm with an outstanding signal-to-noise ratio. Conformational deviations of individual membrane proteins can be observed and their functional states directly imaged. Time-lapse AFM can image membrane proteins at work. In conjunction with high- resolution imaging, the use of the AFM as a single-molecule force spectroscope (SMFS) has gained tremendous importance in recent years. This combination allows to locate the inter- and intramolecular interactions of single membrane proteins. SMFS allows characterization of interactions that guide the folding of proteins and describe the parameters that lead to their destabilization, malfunction and misfolding. Moreover, it enables to measure the interactions established by ligand- and inhibitor-binding and in membrane protein assemblies. Because of its practical use in characterizing various parameters of membrane proteins in their native environment, AFM can be aptly described as a ‘lab on a tip’ device.

Keywords

Atomic Force Microscope Membrane Protein Energy Landscape Atomic Force Microscope Probe Structural Segment 
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.

References

  1. Agre, P., and Kozono, D. (2003). Aquaporin water channels: molecular mechanisms for human diseases. FEBS Lett 555, 72–78.Google Scholar
  2. Alberts, B., Johnson, A., Lewis, J., Raff, M., Roberts, K., and Walter, P. (2002). Molecular Biology of the Cell, 4th ed. (New York, Garland Science).Google Scholar
  3. Alcala, J., Lieska, N., and Maisel, H. (1975). Protein composition of bovine lens cortical fiber cell membranes. Exp Eye Res 21, 581–595.Google Scholar
  4. Andley, U. (2007). Crystallins in the eye: function and pathology. Prog Retin Eye Res 25, 78–98.Google Scholar
  5. Ando, T., Kodera, N., Naito, Y., Kinoshita, T., Furuta, K., and Toyoshima, Y. Y. (2003). A high-speed atomic force microscope for studying biological macromolecules in action. Chemphyschem 4, 1196–1202.Google Scholar
  6. Ando, T., Uchihashi, T., Kodera, N., Yamamoto, D., Taniguch, M., Miyagi, A., and Yamashita, H. (2008). Invited review: high-speed AFM and nano-visualization of biomolecular processes. Eur J Physiol 456, 211–225.Google Scholar
  7. Ando, T., Uchihashi, T., Kodera, N., Yamamoto, D., Taniguchi, M., Miyagi, A., and Yamashita, H. (2007). High-speed atomic force microscopy for observing dynamic biomolecular processes. J Mol Rec 20, 448–458.Google Scholar
  8. Aridor, M., and Hannan, M. L. (2000). Traffic jam: a compendium of human diseases that affect intracellular transport processes. Traffic 1, 836–851.Google Scholar
  9. Aridor, M., and Hannan, M. L. (2002). Traffic jams II: an update of diseases of intracellular transport. Traffic 3, 781–790.Google Scholar
  10. Bell, G. I. (1978). Models for the specific adhesion of cells to cells. Science 200, 618–627.ADSGoogle Scholar
  11. Berman, H. M., Westbrook, J., Feng, Z., Gilliland, G., Bhat, T. N., Weissig, H., Shindyalov, I. N., and Bourne, P. E. (2000). The Protein Data Bank. Nucleic Acids Res 28, 235–242.Google Scholar
  12. Best, R. B., Li, B., Steward, A., Daggett, V., and Clarke, J. (2001). Can non-mechanical proteins withstand force? Stretching barnase by atomic force microscopy and molecular dynamics simulation. Biophys J 81, 2344–2356.Google Scholar
  13. Binnig, G., Quate, C. F., and Gerber, C. (1986). Atomic force microscope. Phys Rev Lett 56, 930–933.ADSGoogle Scholar
  14. Binnig, G., Rohrer, H., Gerber, C., and Weibel, E. (1982). Tunneling through a controllable vacuum gap. Appl Phys Lett 40, 178.ADSGoogle Scholar
  15. Bondar, A. N., Elstner, M., Suhai, S., Smith, J. C., and Fischer, S. (2004). Mechanism of primary proton transfer in bacteriorhodopsin. Structure 12, 1281–1288.Google Scholar
  16. Booth, P. J. (2000). Unravelling the folding of bacteriorhodopsin. Biochim Biophys Acta 1460, 4–14.Google Scholar
  17. Booth, P. J. (2003). The trials and tribulations of membrane protein folding in vitro. Biochim Biophys Acta 1610, 51–56.Google Scholar
  18. Booth, P. J., and Curran, A. R. (1999). Membrane protein folding. Curr Opin Struct Biol 9, 115–121.Google Scholar
  19. Borgia, A., Williams, P. M., and Clarke, J. (2008). Single-molecule studies of protein folding. Annu Rev Biochem 77, 101–125.Google Scholar
  20. Bowie, J. U. (2001). Stabilizing membrane proteins. Curr Opin Struct Biol 11, 397–402.Google Scholar
  21. Breyton, C., Haase, W., Rapoport, T. A., Kuhlbrandt, W., and Collinson, I. (2002). Three-dimensional structure of the bacterial protein-translocation complex SecYEG. Nature 418, 662–665.ADSGoogle Scholar
  22. Brockwell, D. J., Beddard, G. S., Clarkson, J., Zinober, R. C., Blake, A. W., Trinick, J., Olmsted, P. D., Smith, D. A., and Radford, S. E. (2002). The effect of core destabilization on the mechanical resistance of I27. Biophys J 83, 458–472.Google Scholar
  23. Brodsky, J. L., and McCracken, A. A. (1999). ER protein quality control and proteasome-mediated protein degradation. Semin Cell Dev Biol 10, 507–513.Google Scholar
  24. Bryngelson, J. D., Onuchic, J. N., Socci, N. D., and Wolynes, P. G. (1995). Funnels, pathways, and the energy landscape of protein folding: a synthesis. Proteins 21, 167–195.Google Scholar
  25. Butt, H.-J., Downing, K. H., and Hansma, P. K. (1990). Imaging the membrane protein bacteriorhodopsin with the atomic force microscope. Biophys J 58, 1473–1480.Google Scholar
  26. Buzhynskyy, N., Girmens, J.-F., Faigle, W., and Scheuring, S. (2007a). Human cataract lens membrane at subnanometer resolution. J Mol Biol 374, 162–169.Google Scholar
  27. Buzhynskyy, N., Hite, R. K., Walz, T., and Scheuring, S. (2007b). The supramolecular architecture of junctional microdomains in native lens membranes. EMBO Rep 8, 51–55.Google Scholar
  28. Buzhynskyy, N., Sens, P., Prima, V., Sturgis, J. N., and Scheuring, S. (2007c). Rows of ATP synthase dimers in native mitochondrial inner membranes. Biophys J 93, 2870–2876.Google Scholar
  29. Carrion-Vazquez, M., Oberhauser, A. F., Fowler, S. B., Marszalek, P. E., Broedel, S. E., Clarke, J., and Fernandez, J. M. (1999). Mechanical and chemical unfolding of a single protein: a comparison. Proc Natl Acad Sci USA 96, 3694–3699.ADSGoogle Scholar
  30. Chia, Carver, B. C. S., Mulhern, J. A., and Bowie, J. H. (2000). Maculatin 1.1, an anti-microbial peptide from the Australian tree frog, Litoria genimaculata—solution structure and biological activity. Eur J Biochem 267, 1894–1908.Google Scholar
  31. Cisneros, D. A., Oberbarnscheidt, L., Pannier, A., Klare, J. P., Helenius, J., Engelhard, M., Oesterhelt, F., and Muller, D. J. (2008). Transducer binding establishes localized interactions to tune sensory rhodopsin II. Structure, 16, 1206–1213.Google Scholar
  32. Cisneros, D. A., Oesterhelt, D., and Müller, D. J. (2005). Probing origins of molecular interactions stabilizing the membrane proteins halorhodopsin and bacteriorhodopsin. Structure 13, 235–242.Google Scholar
  33. Cotton, R. G., and Horaitis, O. (2002). The HUGO Mutation Database Initiative. Human Genome Organization. Pharmacogenomics J 2, 16–19.Google Scholar
  34. Curnow, P., and Booth, P. J. (2007). Combined kinetic and thermodynamic analysis of alpha-helical membrane protein unfolding. Proc Natl Acad Sci USA 104, 18970–18975.ADSGoogle Scholar
  35. Czajkowsky, D. M., Hotze, E. M., Shao, Z., and Tweten, R. K. (2004). Vertical collapse of a cytolysin prepore moves its transmembrane beta-hairpins to the membrane. EMBO J 23, 3206–3215.Google Scholar
  36. Dill, K. A., and Chan, H. S. (1997). From Levinthal to pathways to funnels. Nat Struct Biol 4, 10–19.Google Scholar
  37. Dobson, C. M. (2003). Protein folding and misfolding. Nature 426, 884–890.ADSGoogle Scholar
  38. Donaldson, P., Kistler, J., and Mathias, R. T. (2001). Molecular solutions to mammalian lens transparency. News Physiol Sci 16, 118–123.Google Scholar
  39. Drake, B., Prater, C. B., Weisenhorn, A. L., Gould, S. A., Albrecht, T. R., Quate, C. F., Cannell, D. S., Hansma, H. G., and Hansma, P. K. (1989). Imaging crystals, polymers, and processes in water with the atomic force microscope. Science 243, 1586–1589.ADSGoogle Scholar
  40. Dupuy, A. D., and Engelman, D. M. (2008). Protein area occupancy at the center of the red blood cell membrane. Proc Natl Acad Sci USA 105, 2848–2852.ADSGoogle Scholar
  41. Elie-Caille, C., Severin, F., Helenius, J., Howard, J., Muller, D. J., and Hyman, A. A. (2007). Straight GDP-tubulin protofilaments form in the presence of taxol. Curr Biol 17, 1765–1770.Google Scholar
  42. Engel, A., and Gaub, H. E. (2008). Structure and mechanics of membrane proteins. Annu Rev Biochem 77, 127–148.Google Scholar
  43. Engel, A., and Müller, D. J. (2000). Observing single biomolecules at work with the atomic force microscope. Nat Struct Biol 7, 715–718.Google Scholar
  44. Engelman, D. M. (2005). Membranes are more mosaic than fluid. Nature 438, 578–580.ADSGoogle Scholar
  45. Engelman, D. M., Chen, Y., Chin, C. N., Curran, A. R., Dixon, A. M., Dupuy, A. D., Lee, A. S., Lehnert, U., Matthews, E. E., Reshetnyak, Y. K., et al. (2003). Membrane protein folding: beyond the two stage model. FEBS Lett 555, 122–125.Google Scholar
  46. Engelman, D. M., and Steitz, T. A. (1981). The spontaneous insertion of proteins into and across membranes: the helical hairpin hypothesis. Cell 23, 411–422.Google Scholar
  47. Evans, E. (1998). Energy landscapes of biomolecular adhesion and receptor anchoring at interfaces explored with dynamic force spectroscopy. Faraday Discuss 111, 1–16.ADSGoogle Scholar
  48. Evans, E. (1999). Looking inside molecular bonds at biological interfaces with dynamic force spectroscopy. Biophys Chem 82, 83–97.Google Scholar
  49. Evans, E. (2001). Probing the relation between force-lifetime and chemistry in single molecular bonds. Annu Rev Biophys Biomol Struct 30, 105–128.Google Scholar
  50. Evans, E., and Ritchie, K. (1997). Dynamic strength of molecular adhesion bonds. Biophys J 72, 1541–1555.Google Scholar
  51. Faham, S., Yang, D., Bare, E., Yohannan, S., Whitelegge, J. P., and Bowie, J.U. (2004). Side-chain contributions to membrane protein structure and stability. J Mol Biol 335, 297–305.Google Scholar
  52. Fandrich, M., Forge, V., Buder, K., Kittler, M., Dobson, C. M., and Diekmann, S. (2003). Myoglobin forms amyloid fibrils by association of unfolded polypeptide segments. Proc Natl Acad Sci USA 100, 15463–15468.ADSGoogle Scholar
  53. Fleschner, C., and Cenedella, R. (1991). Lipid composition of lens plasma membrane fractions enriched in fiber junctions. J Lipid Res 32, 45–53.Google Scholar
  54. Fotiadis, D., Jastrzebska, B., Philippsen, A., Muller, D. J., Palczewski, K., and Engel, A. (2006). Structure of the rhodopsin dimer: a working model for G-protein–coupled receptors. Curr Opin Struct Biol 16, 252–259.Google Scholar
  55. Fotiadis, D., Liang, Y., Filipek, S., Saperstein, D. A., Engel, A., and Palczewski, K. (2003). Atomic-force microscopy: rhodopsin dimers in native disc membranes. Nature 421, 127–128.ADSGoogle Scholar
  56. Frauenfelder, H., Sligar, S. G., and Wolynes, P. G. (1991). The energy landscapes and motions of proteins. Science 254, 1598–1603.ADSGoogle Scholar
  57. Gerber, C., and Lang, H. P. (2006). How the doors to the nanoworld were opened. Nat Nanotechnol 1, 3–5.ADSGoogle Scholar
  58. Gonçalves, R. P., Bernadac, A., Sturgis, J. N., and Scheuring, S. (2005). Architecture of the native photosynthetic apparatus of Phaeospirillum molischianum. J Struct Biol 152, 221–228.Google Scholar
  59. Gonçalves, R. P., Buzhynskyy, N., Prima, V., Sturgis, J. N., and Scheuring, S. (2007). Supramolecular assembly of VDAC in native mitochondrial outer membranes. J Mol Biol 369, 413–418.Google Scholar
  60. Guthold, M., Zhu, X., Rivetti, C., Yang, G., Thomson, N. H., Kasas, S., Hansma, H. G., Smith, B., Hansma, P. K., and Bustamante, C. (1999). Direct observation of one-dimensional diffusion and transcription by Escherichia coli RNA polymerase. Biophys J 77, 2284–2294.Google Scholar
  61. Hansma, H. G., and Laney, D. E. (1996). DNA binding to mica correlates with cationic radius: assay by atomic force microscopy. Biophys J 70, 1933–1939.Google Scholar
  62. Hansma, P. K., Cleveland, J. P., Radmacher, M., Walters, D. A., Hillner, P. E., Bezanilla, M., Fritz, M., Vie, D., Hansma, H. G., Prater, C. B., et al. (1994). Tapping mode atomic force microscopy in liquids. Appl Phys Lett 64, 1738–1740.ADSGoogle Scholar
  63. Hinterdorfer, P., Baumgartner, W., Gruber, H. J., Schilcher, K., and Schindler, H. (1996). Detection and localization of individual antibody–antigen recognition events by atomic force microscopy. Proc Natl Acad Sci USA 93, 3477–3481.ADSGoogle Scholar
  64. Humphris, A. D., Miles, M., and Hobbs, J. K. (2005). A mechanical microscope: high-speed atomic force microscopy. Appl Phys Lett 86, 34106–34109.Google Scholar
  65. Hunt, J. F., Earnest, T. N., Bousche, O., Kalghatgi, K., Reilly, K., Horvath, C., Rothschild, K. J., and Engelman, D. M. (1997a). A biophysical study of integral membrane protein folding. Biochemistry 36, 15156–15176.Google Scholar
  66. Hunt, J. F., Rath, P., Rothschild, K. J., and Engelman, D. M. (1997b). Spontaneous, pH-dependent membrane insertion of a transbilayer alpha-helix. Biochemistry 36, 15177–15192.Google Scholar
  67. Hunte, C., Screpanti, E., Venturi, M., Rimon, A., Padan, E., and Michel, H. (2005). Structure of a Na+/H+ antiporter and insights into mechanism of action and regulation by pH. Nature 435, 1197–1202.ADSGoogle Scholar
  68. Hyeon, C., and Thirumalai, D. (2003). Can energy landscape roughness of proteins and RNA be measured by using mechanical unfolding experiments? Proc Natl Acad Sci USA 100, 10249–10253.ADSGoogle Scholar
  69. Jacobs, D. J., Rader, A. J., Kuhn, L. A., and Thorpe, M. F. (2001). Protein flexibility predictions using graph theory. Proteins 44, 150–165.Google Scholar
  70. Janovjak, H., Kedrov, A., Cisneros, D. A., Sapra, K. T., and Müller, D. J. (2006). Imaging and detecting molecular interactions of single transmembrane proteins. Neurobiol Aging 27, 546–561.Google Scholar
  71. Janovjak, H., Kessler, M., Oesterhelt, D., Gaub, H. E., and Müller, D. J. (2003). Unfolding pathways of native bacteriorhodopsin depend on temperature. EMBO J 22, 5220–5229.Google Scholar
  72. Janovjak, H., Knaus, H., and Muller, D. J. (2007). Transmembrane helices have rough energy surfaces. J Am Chem Soc 129, 246–247.Google Scholar
  73. Janovjak, H., Struckmeier, J., Hubain, M., Kedrov, A., Kessler, M., and Müller, D. J. (2004). Probing the energy landscape of the membrane protein bacteriorhodopsin. Structure 12, 871–879.Google Scholar
  74. Jaroslawski, S., Zadek, B., Ashcroft, F., Venien-Bryan, C., and Scheuring, S. (2007a). Direct visualization of KirBac3.1 potassium channel gating by atomic force microscopy. J Mol Biol 374, 500–505.Google Scholar
  75. Jaroslawski, S., Zadek, B., Ashcroft, F. M., Venien-Bryan, C., and Scheuring, S. (2007b). Direct visualization of KirBac3.1 potassium channel gating by atomic force microscopy. J Mol Biol 374, 500–505.Google Scholar
  76. Joh, N. H., Min, A., Faham, S., Whitelegge, J. P., Yang, D., Woods, V. L., and Bowie, J. U. (2008). Modest stabilization by most hydrogen-bonded side-chain interactions in membrane proteins. Nature 453, 1266–1270.Google Scholar
  77. Kedrov, A., Appel, M., Baumann, H., Ziegler, C., and Muller, D. J. (2008). Examining the dynamic energy landscape of an antiporter upon inhibitor binding. J Mol Biol 375, 1258–1266.Google Scholar
  78. Kedrov, A., Janovjak, H., Sapra, K. T., and Muller, D. J. (2007). Deciphering molecular interactions of native membrane proteins by single-molecule force spectroscopy. Annu Rev Biophys Biomol Struct 36, 233–260.Google Scholar
  79. Kedrov, A., Janovjak, H., Ziegler, C., Kühlbrandt, W., and Müller, D. J. (2006a). Observing folding kinetics and pathways of single antiporters. J Mol Biol 355, 2–8.Google Scholar
  80. Kedrov, A., Krieg, M., Ziegler, C., Kuhlbrandt, W., and Müller, D. J. (2005). Locating ligand binding and activation of a single antiporter. EMBO Rep 6, 668–674.Google Scholar
  81. Kedrov, A., Ziegler, C., Janovjak, H., Kuhlbrandt, W., and Müller, D. J. (2004). Controlled unfolding and refolding of a single sodium-proton antiporter using atomic force microscopy. J Mol Biol 340, 1143–1152.Google Scholar
  82. Kedrov, A., Ziegler, C., and Müller, D. J. (2006b). Differentiating ligand and inhibitor interactions of a single antiporter. J Mol Biol 362, 925–932.Google Scholar
  83. Kessler, M., and Gaub, H. (2006). Unfolding barriers in bacteriorhodopsin probed from the cytoplasmic and the extracellular side by AFM. Structure 14, 521–527.Google Scholar
  84. Kessler, M., Gottschalk, K. E., Janovjak, H., Muller, D. J., and Gaub, H. E. (2006). Bacteriorhodopsin folds into the membrane against an external force. J Mol Biol 357, 644–654.Google Scholar
  85. Khutorsky, V. (2003). Alpha-hairpin stability and folding of transmembrane segments. Biochem Biophys Res Commun 301, 31–34.Google Scholar
  86. Kihara, A., Akiyama, Y., and Ito, K. (1999). Dislocation of membrane proteins in FtsH-mediated proteolysis. EMBO J 18, 2970–2981.Google Scholar
  87. Klein-Seetharaman, J. (2005). Dual role of interactions between membranous and soluble portions of helical membrane receptors for folding and signaling. Trends Pharmacol Sci 26, 183–189.Google Scholar
  88. Klyszejko, A. L., Shastri, S., Mari, S. A., Grubmuller, H., Muller, D. J., and Glaubitz, C. (2008). Folding and assembly of proteorhodopsin. J Mol Biol 376, 35–41.Google Scholar
  89. Kobayashi, S., Takeshima, K., Park, C. B., Kim, S. C., and Matsuzaki, K. (2000). Interactions of the novel antimicrobial peptide buforin 2 with lipid bilayers: proline as a translocation promoting factor. Biochemistry 39, 8648–8654.Google Scholar
  90. Kopito, R. R. (1999). Biosynthesis and degradation of CFTR. Physiol Rev 79, S167–S173.Google Scholar
  91. Ladokhin, A. S., and White, S. H. (1999). Folding of amphipathic alpha-helices on membranes: energetics of helix formation by melittin. J Mol Biol 285, 1363–1369.Google Scholar
  92. Ladokhin, A. S., and White, S. H. (2004). Interfacial folding and membrane insertion of a designed helical peptide. Biochemistry 43, 5782–5791.Google Scholar
  93. Lanyi, J. K. (1997). Mechanisms of ion transport across membranes. J Biol Chem 272, 31209–31212.Google Scholar
  94. Lazarova, T., Sanz, C., Querol, E., and Padros, E. (2000). Fourier transform infrared evidence for early deprotonation of Asp85 at alkaline pH in the photocycle of bacteriorhodopsin mutants containing E194Q. Biophys J 78, 2022–2030.Google Scholar
  95. Leonhard, K., Guiard, B., Pellecchia, G., Tzagoloff, A., Neupert, W., and Langer, T. (2000). Membrane protein degradation by AAA proteases in mitochondria: extraction of substrates from either membrane surface. Mol Cell 5, 629–638.Google Scholar
  96. Li, H., Carrion-Vazquez, M., Oberhauser, A. F., Marszalek, P. E., and Fernandez, J. M. (2000). Point mutations alter the mechanical stability of immunoglobulin modules. Nat Struct Biol 7, 1117–1120.Google Scholar
  97. Lu, H., Marti, T., and Booth, P. J. (2001). Proline residues in transmembrane alpha helices affect the folding of bacteriorhodopsin. J Mol Biol 308, 437–446.Google Scholar
  98. Luecke, H., Richter, H.-T., and Lanyi, J. K. (1998). Proton transfer pathways in bacteriorhodopsin at 2.3 Angstrom resolution. Science 280, 1934–1937.ADSGoogle Scholar
  99. Mangenot, S., Buzhynskyy, N., Girmens, J.-F., and Scheuring, S. (2008). Malformation of junctional microdomains in type II diabetic cataract lens membranes. pflugers Arch 457, 1265–1274.Google Scholar
  100. McKibbin, C., Farmer, N. A., Jeans, C., Reeves, P. J., Khorana, H. G., Wallace, B. A., Edwards, P. C., Villa, C., and Booth, P. J. (2007). Opsin stability and folding: modulation by phospholipid bicelles. J Mol Biol 374, 1319–1332.Google Scholar
  101. Mitsuoka, K., Hirai, T., Murata, K., Miyazawa, A., Kidera, A., Kimura, Y., and Fujiyoshi, Y. (1999). The structure of bacteriorhodopsin at 3.0 Å resolution based on electron crystallography: implication of the charge distribution. J Mol Biol 286, 861–882.Google Scholar
  102. Möller, C., Allen, M., Elings, V., Engel, A., and Müller, D. J. (1999). Tapping-mode atomic force microscopy produces faithful high-resolution images of protein surfaces. Biophys J 77, 1150–1158.Google Scholar
  103. Möller, C., Fotiadis, D., Suda, K., Engel, A., Kessler, M., and Müller, D. J. (2003). Determining molecular forces that stabilize human aquaporin-1. J Struct Biol 142, 369–378.Google Scholar
  104. Müller, D. J., Amrein, M., and Engel, A. (1997). Adsorption of biological molecules to a solid support for scanning probe microscopy. J Struct Biol 119, 172–188.Google Scholar
  105. Müller, D. J., Baumeister, W., and Engel, A. (1996). Conformational change of the hexagonally packed intermediate layer of Deinococcus radiodurans monitored by atomic force microscopy. J Bacteriol 178, 3025–3030.Google Scholar
  106. Muller, D. J., and Dufrene, Y. F. (2008). Atomic force microscopy as a multifunctional molecular toolbox in nanobiotechnology. Nat Nanotechnol 3, 261–269.ADSGoogle Scholar
  107. Muller, D. J., and Engel, A. (2007). Atomic force microscopy and spectroscopy of native membrane proteins. Nature Protocols 2, 2191–2197.Google Scholar
  108. Müller, D. J., and Engel, A. (1999). pH and voltage induced structural changes of porin OmpF explain channel closure. J Mol Biol 285, 1347–1351.Google Scholar
  109. Müller, D. J., Engel, A., Matthey, U., Meier, T., Dimroth, P., and Suda, K. (2003). Observing membrane protein diffusion at subnanometer resolution. J Mol Biol 327, 925–930.Google Scholar
  110. Müller, D. J., Fotiadis, D., Scheuring, S., Müller, S. A., and Engel, A. (1999). Electrostatically balanced subnanometer imaging of biological specimens by atomic force microscopy. Biophys J 76, 1101–1111.Google Scholar
  111. Müller, D. J., Hand, G. M., Engel, A., and Sosinsky, G. (2002). Conformational changes in surface structures of isolated connexin26 gap junctions. EMBO J 21, 3598–3607.Google Scholar
  112. Müller, D. J., Kessler, M., Oesterhelt, F., Möller, C., Oesterhelt, D., and Gaub, H. (2002c). Stability of bacteriorhodopsin alpha-helices and loops analyzed by single-molecule force spectroscopy. Biophys J 83, 3578–3588.Google Scholar
  113. Muller, D. J., Sapra, K. T., Scheuring, S., Kedrov, A., Frederix, P. L., Fotiadis, D., and Engel, A. (2006). Single-molecule studies of membrane proteins. Curr Opin Struct Biol 16, 489–495.Google Scholar
  114. Neuman, K. C., and Nagy, A. (2008). Single-molecule force spectroscopy: optical tweezers, magnetic tweezers and atomic force microscopy. Nat Methods 5, 491–505.Google Scholar
  115. Oberhauser, A. F., Marszalek, P. E., Carrion-Vazquez, M., and Fernandez, J. M. (1999). Single protein misfolding events captured by atomic force microscopy. Nat Struct Biol 6, 1025–1028.Google Scholar
  116. Oesterhelt, F., Oesterhelt, D., Pfeiffer, M., Engel, A., Gaub, H. E., and Müller, D. J. (2000). Unfolding pathways of individual bacteriorhodopsins. Science 288, 143–146.ADSGoogle Scholar
  117. Okada, T., Sugihara, M., Bondar, A. N., Elstner, M., Entel, P., and Buss, V. (2004). The retinal conformation and its environment in rhodopsin in light of a new 2.2 A crystal structure. J Mol Biol 342, 571–583.Google Scholar
  118. Oliveberg, M., and Wolynes, P.G. (2006). The experimental survey of protein–folding energy landscapes. Q Rev Biophys, 1–44.Google Scholar
  119. Overington, J. P., Al-Lazikani, B., and Hopkins, A. L. (2006). How many drug targets are there? Nat Rev Drug Discov 5, 993–996.Google Scholar
  120. Padan, E., Venturi, M., Gerchman, Y., and Dover, N. (2001). Na(+)/H(+) antiporters. Biochim Biophys Acta 1505, 144–157.Google Scholar
  121. Park, P. S., Sapra, K. T., Kolinski, M., Filipek, S., Palczewski, K., and Muller, D. J. (2007). Stabilizing effect of Zn2+ in native bovine rhodopsin. J Biol Chem 282, 11377–11385.Google Scholar
  122. Park, P. S.-H., and Palczewski, K. (2005). Diversifying the repertoire of G protein–coupled receptors through oligomerization. Proc Natl Acad Sci USA 102, 8793–8794.ADSGoogle Scholar
  123. Popot, J. L., and Engelman, D. M. (1990). Membrane protein folding and oligomerization: the two-stage model. Biochemistry 29, 4031–4037.Google Scholar
  124. Rader, A. J., Anderson, G., Isin, B., Khorana, H. G., Bahar, I., and Klein-Seetharaman, J. (2004). Identification of core amino acids stabilizing rhodopsin. Proc Natl Acad Sci USA 101, 7246–7251.ADSGoogle Scholar
  125. Rajendran, L., and Simons, K. (2005). Lipid rafts and membrane dynamics. J Cell Sci 118, 1099–1102.Google Scholar
  126. Rapoport, T. A. (2007). Protein translocation across the eukaryotic endoplasmic reticulum and bacterial plasma membranes. Nature 450, 663–669.ADSGoogle Scholar
  127. Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J. M., and Gaub, H. E. (1997). Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 276, 1109–1112.Google Scholar
  128. Riley, M. L., Wallace, B. A., Flitsch, S. L., and Booth, P. J. (1997). Slow alpha helix formation during folding of a membrane protein. Biochemistry 36, 192–196.Google Scholar
  129. Sadlish, H., Pitonzo, D., Johnson, A. E., and Skach, W. R. (2005). Sequential triage of transmembrane segments by Sec61alpha during biogenesis of a native multispanning membrane protein. Nat Struct Mol Biol 12, 870–878.Google Scholar
  130. Sahin, O., Magonov, S., Su, C., Quate, C., and Solgaard, O. (2007). An atomic force microscope tip designed to measure time-varying nanomechanical forces. Nat Nanotechnol 2, 507–514.Google Scholar
  131. Sanders, C. R., and Myers, J. K. (2004). Disease-related misassembly of membrane proteins. Annu Rev Biophys Biomol Struct 33, 25–51.Google Scholar
  132. Sanders, C. R., and Nagy, J. K. (2000). Misfolding of membrane proteins in health and disease: the lady or the tiger? Curr Opin Struct Biol 10, 438–442.Google Scholar
  133. Sanz, C., Marquez, M., Peralvarez, A., Elouatik, S., Sepulcre, F., Querol, E., Lazarova, T., and Padros, E. (2001). Contribution of extracellular Glu residues to the structure and function of bacteriorhodopsin. Presence of specific cation-binding sites. J Biol Chem 276, 40788–40794.Google Scholar
  134. Sapra, K. T., Balasubramanian, G. P., Labudde, D., Bowie, J. U., and Muller, D. J. (2008a). Point mutations in membrane proteins reshape energy landscape and populate different unfolding pathways. J Mol Biol 376, 1076–1090.Google Scholar
  135. Sapra, K. T., Besir, H., Oesterhelt, D., and Muller, D. J. (2006a). Characterizing molecular interactions in different bacteriorhodopsin assemblies by single-molecule force spectroscopy. J Mol Biol 355, 640–650.Google Scholar
  136. Sapra, K. T., Doehner, J., Renugopalakrishnan, V., Padros, E., and Muller, D. J. (2008b). Role of extracellular glutamic acids in the stability and energy landscape of bacteriorhodopsin. Biophys J, 95, 3407–3418.Google Scholar
  137. Sapra, K. T., Park, P. S., Filipek, S., Engel, A., Müller, D. J., and Palczewski, K. (2006b). Detecting molecular interactions that stabilize native bovine rhodopsin. J Mol Biol 358, 255–269.Google Scholar
  138. Sapra, K. T., Park, P. S., Palczewski, K., and Muller, D. J. (2008c). Mechanical properties of bovine rhodopsin and bacteriorhodopsin: possible roles in folding and function. Langmuir 24, 1330–1337.Google Scholar
  139. Scheuring, S. (2006). AFM studies of the supramolecular assembly of bacterial photosynthetic core-complexes. Curr Opinion Chem Biol 10, 387–393.Google Scholar
  140. Scheuring, S., Boudier, T., and Sturgis, J. N. (2007a). From high-resolution AFM topographs to atomic models of supramolecular assemblies. J Struct Biol 159, 268–276.Google Scholar
  141. Scheuring, S., Buzhynskyy, N., Jaroslawski, S., Gonçalves, R. P., Hite, R. K., and Walz, T. (2007b). Structural models of the supramolecular organization of AQP0 and connexons in junctional microdomains J Struct Biol 160, 385–394.Google Scholar
  142. Scheuring, S., Buzhysnskyy, N., Gonçalves, R., and Jaroslawski, S. (2007c). Atomic force microscopy: high-resolution imaging of structure and assembly of membrane proteins. In Biophysical Analysis of Membrane Proteins, E. Pebay-Peyroula, ed. (Wiley, Chichester, UK) 141–158.Google Scholar
  143. Scheuring, S., Fotiadis, D., Möller, C., Müller, S. A., Engel, A., and Müller, D. J. (2001). Single proteins observed by atomic force microscopy. Single Molecules 2, 59–67.ADSGoogle Scholar
  144. Scheuring, S., Gonçalves, R. P., Prima, V., and Sturgis, J. N. (2006). The photosynthetic apparatus of Rhodopseudomonas palustris: structures and organization. J Mol Biol 358, 83–96.Google Scholar
  145. Scheuring, S., Levy, D., and Rigaud, J.-L. (2005). Watching the components of photosynthetic bacterial membranes and their “in situ” organization by atomic force microscopy. Biochim Biophys Acta 1712, 109–127.Google Scholar
  146. Scheuring, S., Rigaud, J.-L., and Sturgis, J. N. (2004a). Variable LH2 stoichiometry and core clustering in native membranes of Rhodospirillum photometricum. EMBO J 23, 4127–4133.Google Scholar
  147. Scheuring, S., Ringler, P., Borgina, M., Stahlberg, H., Müller, D. J., Agre, P., and Engel, A. (1999). High resolution topographs of the Escherichia coli waterchannel aquaporin Z. EMBO J 18, 4981–4987.Google Scholar
  148. Scheuring, S., and Sturgis, J. N. (2005). Chromatic adaptation of photosynthetic membranes. Science 309, 484–487.ADSGoogle Scholar
  149. Scheuring, S., and Sturgis, J. N. (2006). Dynamics and diffusion in photosynthetic membranes from Rhodospirillum photometricum. Biophys J 91, 3707–3717.Google Scholar
  150. Scheuring, S., Sturgis, J. N., Prima, V., Bernadac, A., Lévy, D., and Rigaud, J.-L. (2004b). Watching the photosynthetic apparatus in native membranes. Proc Natl Acad Sci USA 101, 11293–11297.ADSGoogle Scholar
  151. Schwesinger, F., Ros, R., Strunz, T., Anselmetti, D., Guntherodt, H. J., Honegger, A., Jermutus, L., Tiefenauer, L., and Pluckthun, A. (2000). Unbinding forces of single antibody–antigen complexes correlate with their thermal dissociation rates. Proc Natl Acad Sci USA 97, 9972–9977.ADSGoogle Scholar
  152. Seddon, A. M., Curnow, P., and Booth, P. J. (2004). Membrane proteins, lipids and detergents: not just a soap opera. Biochim Biophys Acta 1666, 105–117.Google Scholar
  153. Seibert, F. S., Loo, T. W., Clarke, D. M., and Riordan, J. R. (1997). Cystic fibrosis: channel, catalytic, and folding properties of the CFTR protein. J Bioenerg Biomembr 29, 429–442.Google Scholar
  154. Singer, S. J., and Nicolson, G. L. (1972). The fluid mosaic model of the structure of cell membranes. Science 175, 720–731.ADSGoogle Scholar
  155. Soekarjo, M., Eisenhawer, M., Kuhn, A., and Vogel, H. (1996). Thermodynamics of the membrane insertion process of the M13 procoat protein, a lipid bilayer traversing protein containing a leader sequence. Biochemistry 35, 1232–1241.Google Scholar
  156. Stahlberg, H., Müller, D. J., Suda, K., Fotiadis, D., Engel, A., Matthey, U., Meier, T., and Dimroth, P. (2001). Bacterial ATP synthase has an undacemeric rotor. EMBO Rep 2, 229–235.Google Scholar
  157. Stoffler, D., Goldie, K. N., Feja, B., and Aebi, U. (1999). Calcium-mediated structural changes of native nuclear pore complexes monitored by time-lapse atomic force microscopy. J Mol Biol 287, 741–752.Google Scholar
  158. Struckmeier, J., Wahl, R., Leuschner, M., Nunes, J., Janovjak, H., Geisler, U., Hofmann, G., Jahnke, T., and Müller, D. J. (2008). Fully automated single-molecule force spectroscopy for screening applications. Nanotechnology, 19, 384020–384030.Google Scholar
  159. Strunz, T., Oroszlan, K., Schafer, R., and Guntherodt, H. J. (1999). Dynamic force spectroscopy of single DNA molecules. Proc Natl Acad Sci USA 96, 11277–11282.ADSGoogle Scholar
  160. Sudo, Y., and Spudich, J. L. (2006). Three strategically placed hydrogen-bonding residues convert a proton pump into a sensory receptor. Proc Natl Acad Sci USA 103, 16129–16134.ADSGoogle Scholar
  161. Taglicht, D., Padan, E., and Schuldiner, S. (1991). Overproduction and purification of a functional Na+/H+ antiporter coded by nhaA (ant) from Escherichia coli. J Biol Chem 266, 11289–11294.Google Scholar
  162. Tastan, O., Yu, E., Ganapathiraju, M., Aref, A., Rader, A. J., and Klein-Seetharaman, J. (2007). Comparison of stability predictions and simulated unfolding of rhodopsin structures. Photochem Photobiol 83, 351–362.Google Scholar
  163. Tsai, B., Ye, Y., and Rapoport, T. A. (2002). Retrotranslocation of proteins from the endoplasmic reticulum into the cytosol. Nat Rev Mol Cell Biol 3, 246–255.Google Scholar
  164. Van den Berg, B., Clemons, W. M. J., Collinson, I., Modis, Y., Hartmann, E., Harrison, S.C., and Rapoport, T. A. (2004). X-ray structure of a protein-conducting channel. Nature 427, 36–44.ADSGoogle Scholar
  165. Vendruscolo, M., and Dobson, C. M. (2005). A glimpse at the organization of the protein universe. Proc Natl Acad Sci USA 102, 5641–5642.ADSGoogle Scholar
  166. Viani, M. B., Pietrasanta, L. I., Thompson, J. B., Chand, A., Gebeshuber, I. C., Kindt, J. H., Richter, M., Hansma, H. G., and Hansma, P. K. (2000). Probing protein–protein interactions in real time. Nat Struct Biol 7, 644–647.Google Scholar
  167. Viani, M. B., Schäffer, T. E., Paloczi, G. T., Pietrasanta, L. I., Smith, B. L., Thompson, J. B., Richter, M., Rief, M., Gaub, H. E., Plaxco, K. W., et al. (1999). Fast imaging and fast force spectroscopy of single biopolymers with a new atomic force microscope designed for small cantilevers. Rev Sci Instrum 70, 4300–4303.ADSGoogle Scholar
  168. White, S. (2008). Membrane proteins of known 3D structure. Available at: http://blanco.biomol.uci.edu/Membrane_Proteins_xtal.html.
  169. Wieprecht, T., Beyermann, M., and Seelig, J. (2002). Thermodynamics of the coil–alpha-helix transition of amphipathic peptides in a membrane environment: the role of vesicle curvature. Biophys Chem 96, 191–201.Google Scholar
  170. Williams, P. M., Fowler, S. B., Best, R. B., Toca-Herrera, J. L., Scott, K. A., Steward, A., and Clarke, J. (2003). Hidden complexity in the mechanical properties of titin. Nature 422, 446–449.ADSGoogle Scholar
  171. Wolynes, P. G., Onuchic, J. N., and Thirumalai, D. (1995). Navigating the folding routes. Science 267, 1619–1620.ADSGoogle Scholar
  172. Xie, K., Hessa, T., Seppala, S., Rapp, M., von Heijne, G., and Dalbey, R. E. (2007). Features of transmembrane segments that promote the lateral release from the translocase into the lipid phase. Biochemistry 46, 15153–15161.Google Scholar
  173. Yamamoto, D., Uchihashi, T., Kodera, N., and Ando, T. (2008). Anisotropic diffusion of point defects in two-dimensional crystal of streptavidin observed by high-speed atomic force microscopy. Nanotechnology, 19, 384009–384018.Google Scholar
  174. Ye, Y., Meyer, H. H., and Rapoport, T. A. (2001). The AAA ATPase Cdc48/p97 and its partners transport proteins from the ER into the cytosol. Nature 414, 652–656.ADSGoogle Scholar
  175. Yohannan, S., Faham, S., Yang, D., Whitelegge, J. P., and Bowie, J. U. (2004). The evolution of transmembrane helix kinks and the structural diversity of G protein–coupled receptors. Proc Natl Acad Sci USA 101, 959–963.ADSGoogle Scholar
  176. Yu, J., Bippes, C. A., Hand, G. M., Muller, D. J., and Sosinsky, G. E. (2007). Aminosulfonate modulated pH-induced conformational changes in connexin26 hemichannels. J Biol Chem 282, 8895–8904.Google Scholar

Copyright information

© Springer Science+Business Media, LLC 2009

Authors and Affiliations

  • S. Scheuring
    • 1
  • K. Tanuj Sapra
    • 1
  • Daniel J. Müller
    • 1
  1. 1.Institut Curie, UMR168-CNRS75248 ParisFrance

Personalised recommendations