SDynamic Force Spectroscopy with the Atomic Force Microscope

  • Phil Williams

The dynamic force spectroscopy (DFS) experiment has been with us for nearly a decade [1]. By studying the effect of force on the dissociation kinetics of molecular interactions, hitherto hidden information about physics, chemistry, and biology is gained. Since the statement of the theory and the first demonstration of the experiment [2], we have seen developments in theory, experimental practice, and data analysis. Advances in theory have suggested the possibility of measuring more than dissociation rates over transition states and their displacements, such as the change in energy of the system at the transition state [3–5], the roughness of the dissociation landscape [6–8], and equilibrium phenomena [9]. Today, the instrumentation used to undertake DFS that is most prevalent in the literature is the atomic force microscope (AFM). The significant advances we have seen in both theory and experiment have sometimes taken place in isolation, and here I believe it is worth considering the application of DFS with current AFM technology. How accurately can we do DFS with an AFM? What exactly can we measure with the AFM, and what advances are needed?


Transition State Dissociation Rate Bond Rupture Force Spectroscopy Rupture Force 
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  1. 1.
    Evans, E. and K. Ritchie, Dynamic strength of molecular adhesion bonds. Biophysical Journal, 1997. 72(4): pp. 1541–1555.CrossRefADSGoogle Scholar
  2. 2.
    Merkel, R., et al, Energy landscapes of receptor-ligand bonds explored with dynamic force spectroscopy. Nature, 1999. 397(6714): pp. 50–53.CrossRefADSGoogle Scholar
  3. 3.
    Hummer, G. and A. Szabo, Kinetics from nonequilibrium single-molecule pulling experiments. Biophysical Journal, 2003. 85(1): pp. 5–15.CrossRefADSGoogle Scholar
  4. 4.
    Hummer, G. and A. Szabo, Free energy surfaces from single-molecule force spectroscopy. Accounts of Chemical Research, 2005. 38(7): pp. 504–513.CrossRefGoogle Scholar
  5. 5.
    Dudko, O.K., G. Hummer, and A. Szabo, Intrinsic rates and activation free energies from single-molecule pulling experiments. Physical Review Letters, 2006. 96(10).Google Scholar
  6. 6.
    Hyeon, C.B. and D. Thirumalai, Can energy landscape roughness of proteins and RNA be measured by using mechanical unfolding experiments? Proceedings of the National Academy of Sciences of the United States of America, 2003. 100(18): pp. 10249–10253.CrossRefADSGoogle Scholar
  7. 7.
    Nevo, R., et al, Direct measurement of protein energy landscape roughness. Embo Reports, 2005. 6(5): pp. 482–486.CrossRefGoogle Scholar
  8. 8.
    Schlierf, M. and M. Rief, Temperature softening of a protein in single-molecule experiments. Journal of Molecular Biology, 2005. 354(2): pp. 497–503.CrossRefGoogle Scholar
  9. 9.
    Evans, E. and P.M. Williams, eds. Dynamic Force Spectroscopy: I. Single Bonds. Les Houches Ecole de Physique LVVX. Physics of Bio-molecules and Cells, ed. H. Flyvbjerg. 2002, Springer.Google Scholar
  10. 10.
    Evans, E., Probing the relation between force - Lifetime - and chemistry in single molecular bonds. Annual Review of Biophysics and Biomolecular Structure, 2001. 30: pp. 105–128.CrossRefGoogle Scholar
  11. 11.
    Evans, E., et al, Chemically distinct transition states govern rapid dissociation of single L-selectin bonds under force. Proceedings of the National Academy of Sciences of the United States of America, 2001. 98(7): pp. 3784–3789.CrossRefADSGoogle Scholar
  12. 12.
    Green, N.M., Avidin. Advances in Protein Chemistry, 1975. 29: p. 85–133.CrossRefGoogle Scholar
  13. 13.
    Green, N.M., Avidin and Streptavidin. Methods in Enzymology, 1990. 184: pp. 51–67.CrossRefGoogle Scholar
  14. 14.
    Green, N.H., et al, Single-molecule investigations of RNA dissociation. Biophysical Journal, 2004. 86(6): p. 3811–3821.CrossRefADSGoogle Scholar
  15. 15.
    Abramowitz, M. and I.A. Stegun, eds. Hanbook of Mathematical Functions. 1972, Dover.Google Scholar
  16. 16.
    Williams, P.M., Analytical descriptions of dynamic force spectroscopy: behaviour of multiple connections. Analytica Chimica Acta, 2003. 479(1): pp. 107–115.CrossRefGoogle Scholar
  17. 17.
    Gergely, C., et al, Unbinding process of adsorbed proteins under external stress studied by atomic force microscopy spectroscopy. Proceedings of the National Academy of Sciences of the United States of America, 2000. 97(20): pp. 10802–10807.CrossRefADSGoogle Scholar
  18. 18.
    Evans, E., et al, Mechanical switching and coupling between two dissociation pathways in a P-selectin adhesion bond. Proceedings of the National Academy of Sciences of the United States of America, 2004. 101(31): pp. 11281–11286.CrossRefADSGoogle Scholar
  19. 19.
    Williams, P.M., et al, On the dynamic behaviour of the forced dissociation of ligand-receptor pairs. Journal of the Chemical Society-Perkin Transactions 2, 2000(1): pp. 5–8.CrossRefGoogle Scholar
  20. 20.
    Lo, Y.S., Y.J. Zhu, and T.P. Beebe, Loading-rate dependence of individual ligand-receptor bond-rupture forces studied by atomic force microscopy. Langmuir, 2001. 17(12): pp. 3741–3748.CrossRefGoogle Scholar
  21. 21.
    Grubmuller, H., B. Heymann, and P. Tavan, Ligand binding: Molecular mechanics calculation of the streptavidin biotin rupture force. Science, 1996. 271(5251): pp. 997–999.CrossRefADSGoogle Scholar
  22. 22.
    Moore, A., et al., Enthalpic approach to the analysis of the scanning force ligand rupture experiment. Journal of the Chemical Society-Perkin Transactions 2, 1998(2): pp. 253–258.CrossRefGoogle Scholar
  23. 23.
    Moore, A., et al., Analyzing the origins of receptor-ligand adhesion forces measured by the scanning force microscope. Journal of the Chemical Society-Perkin Transactions 2, 1999(3): pp. 419–423.CrossRefGoogle Scholar
  24. 24.
    Galligan, E., et al, Simulating the dynamic strength of molecular interactions. Journal of Chemical Physics, 2001. 114(7): pp. 3208–3214.CrossRefADSGoogle Scholar
  25. 25.
    Williams, P.M., Force Spectroscopy, in Scanning Probe Microscopies: Beyond Imaging, P. Samori, Editor. 2006, WILEY-VCH. pp. 250–274.Google Scholar
  26. 26.
    Tawar, R.G., et al., Complex bond architecture revealed through dynamic force spectroscopy.Google Scholar
  27. 27.
    Ros, R., et al, Antigen binding forces of individually addressed single-chain Fv antibody molecules. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(13): pp. 7402–7405.CrossRefADSGoogle Scholar
  28. 28.
    Patel, A.B., et al, Influence of architecture on the kinetic stability of molecular assemblies. Journal of the American Chemical Society, 2004. 126(5): pp. 1318–1319.CrossRefGoogle Scholar
  29. 29.
    Sulchek, T.A., et al, Dynamic force spectroscopy of parallel individual Mucin1-antibody bonds. Proceedings of the National Academy of Sciences of the United States of America, 2005. 102(46): pp. 16638–16643.CrossRefADSGoogle Scholar
  30. 30.
    Sulchek, T., R.W. Friddle, and A. Noy, Strength of multiple parallel biological bonds. Biophysical Journal, 2006. 90(12): pp. 4686–4691.CrossRefADSGoogle Scholar
  31. 31.
    Cleveland, J.P., et al, A Nondestructive Method for Determining the Spring Constant of Cantilevers for Scanning Force Microscopy. Review of Scientific Instruments, 1993. 64(2): pp. 403–405.CrossRefADSGoogle Scholar
  32. 32.
    Walters, D.A., et al, Short cantilevers for atomic force microscopy. Review of Scientific Instruments, 1996. 67(10): pp. 3583–3590.CrossRefADSGoogle Scholar
  33. 33.
    Heim, L.O., M. Kappl, and H.J. Butt, Tilt of atomic force microscope cantilevers: Effect on spring constant and adhesion measurements. Langmuir, 2004. 20(7): pp. 2760–2764.CrossRefGoogle Scholar
  34. 34.
    Proksch, R., et al, Finite optical spot size and position corrections in thermal spring constant calibration. Nanotechnology, 2004. 15(9): pp. 1344–1350.CrossRefADSGoogle Scholar
  35. 35.
    Bonaccurso, E. and H.J. Butt, Microdrops on atomic force microscope cantilevers: Evaporation of water and spring constant calibration. Journal of Physical Chemistry B, 2005. 109(1): pp. 253–263.CrossRefGoogle Scholar
  36. 36.
    Clifford, C.A. and M.P. Seah, The determination of atomic force microscope cantilever spring constants via dimensional methods for nanomechanical analysis. Nanotechnology, 2005. 16(9): pp. 1666–1680.CrossRefADSGoogle Scholar
  37. 37.
    Stiernstedt, J., M.W. Rutland, and P. Attard, A novel technique for the in situ calibration and measurement of friction with the atomic force microscope. Review of Scientific Instruments, 2005. 76(8).Google Scholar
  38. 38.
    Higgins, M.J., et al., Noninvasive determination of optical lever sensitivity in atomic force microscopy. Review of Scientific Instruments, 2006. 77(1).Google Scholar
  39. 39.
    Ng, S.P., et al, Mechanical unfolding of TNfn3: The unfolding pathway of a fnIII domain probed by protein engineering, AFM and MD simulation. Journal of Molecular Biology, 2005. 350(4): pp. 776–789.CrossRefGoogle Scholar
  40. 40.
    Wojcikiewicz, E.P., et al, Force spectroscopy of LFA-1 and its ligands, ICAM-1 and ICAM-2. Biomacromolecules, 2006. 7(11): pp. 3188–3195.CrossRefGoogle Scholar
  41. 41.
    Evans, E. and K. Ritchie, Strength of a weak bond connecting flexible polymer chains. Biophysical Journal, 1999. 76(5): pp. 2439–2447.CrossRefADSGoogle Scholar
  42. 42.
    Sturges, H., The choice of a class-interval. Journal of the American Statistical Association, 1926. 21: pp. 65–66.Google Scholar
  43. 43.
    Scott, D., On optimal and data-based histograms. Biometrika, 1979. 66(3): pp. 605–610.zbMATHCrossRefMathSciNetGoogle Scholar
  44. 44.
    Baumgartner, W., et al, Cadherin interaction probed by atomic force microscopy. Proceedings of the National Academy of Sciences of the United States of America, 2000. 97(8): pp. 4005–4010.CrossRefADSGoogle Scholar
  45. 45.
    Clarke, J. and P.M. Williams, Unfolding Induced by Mechanical Force, in Protein Folding Handbook, J. Buchner and T. Kiefhaber, Editors. 2005, WILEY-VCH. pp. 1111–1142.CrossRefGoogle Scholar
  46. 46.
    Wahab, O., et al., Multiple transition states measured in RNA strand dissociation.Google Scholar
  47. 47.
    Tees, D.F.J., R.E. Waugh, and D.A. Hammer, A microcantilever device to assess the effect of force on the lifetime of selectin-carbohydrate bonds. Biophysical Journal, 2001. 80(2): pp. 668–682.CrossRefADSGoogle Scholar
  48. 48.
    Clarke, R.J., et al, The drag on a microcantilever oscillating near a wall. Journal of Fluid Mechanics, 2005. 545: pp. 397–426.zbMATHCrossRefADSMathSciNetGoogle Scholar
  49. 49.
    Clarke, R.J., et al., Stochastic elastohydrodynamics of a microcantilever oscillating near a wall. Physical Review Letters, 2006. 96(5).Google Scholar
  50. 50.
    Clarke, R.J., et al, Three-dimensional flow due to a microcantilever oscillating near a wall: an unsteady slender-body analysis. Proceedings of the Royal Society a-Mathematical Physical and Engineering Sciences, 2006. 462(2067): pp. 913–933.zbMATHCrossRefADSGoogle Scholar
  51. 51.
    Janovjak, H.J., J. Struckmeier, and D.J. Muller, Hydrodynamic effects in fast AFM single-molecule force measurements. European Biophysics Journal with Biophysics Letters, 2005. 34(1): pp. 91–96.Google Scholar
  52. 52.
    Best, R.B. and J. Clarke, What can atomic force microscopy tell us about protein folding? Chemical Communications, 2002(3): pp. 183–192.Google Scholar
  53. 53.
    Best, R.B., et al, Force mode atomic force microscopy as a tool for protein folding studies. Analytica Chimica Acta, 2003. 479(1): pp. 87–105.CrossRefGoogle Scholar
  54. 54.
    Clarke, J. and G. Schreiber, Folding and binding - new technologies and new perspectives - Editorial overview. Current Opinion in Structural Biology, 2003. 13(1): pp. 71–74.CrossRefGoogle Scholar
  55. 55.
    Carrion-Vazquez, M., et al, AFM and chemical unfolding of a single protein follow the same pathway. Biophysical Journal, 1999. 76(1): pp. A173-A173.Google Scholar
  56. 56.
    Carrion-Vazquez, M., et al, Mechanical and chemical unfolding of a single protein: A comparison. Proceedings of the National Academy of Sciences of the United States of America, 1999. 96(7): pp. 3694–3699.CrossRefADSGoogle Scholar
  57. 57.
    Fowler, S.B., et al, Mechanical unfolding of a titin Ig domain: Structure of unfolding intermediate revealed by combining AFM, molecular dynamics simulations, NMR and protein engineering. Journal of Molecular Biology, 2002. 322(4): pp. 841–849.CrossRefMathSciNetGoogle Scholar
  58. 58.
    Williams, P.M., et al, Hidden complexity in the mechanical properties of titin. Nature, 2003. 422(6930): pp. 446–449.CrossRefADSGoogle Scholar
  59. 59.
    Lu, H., et al, Unfolding of titin immunoglobulin domains by steered molecular dynamics simulation. Biophysical Journal, 1998. 75(2): pp. 662–671.CrossRefADSGoogle Scholar
  60. 60.
    Marszalek, P.E., et al, Mechanical unfolding intermediates in titin modules. Nature, 1999. 402(6757): pp. 100–103.CrossRefADSGoogle Scholar
  61. 61.
    Williams, P.M. and E. Evans, eds. Dynamic Force Spectroscopy: II. Multiple Bonds. Les Houches Ecole de Physique LVVX. Physics of Bio-molecules and Cells, ed. H. Flyvbjerg. 2002, Springer.Google Scholar
  62. 62.
    Zinober, R.C., et al, Mechanically unfolding proteins: The effect of unfolding history and the supramolecular scaffold. Protein Science, 2002. 11(12): pp. 2759–2765.CrossRefGoogle Scholar
  63. 63.
    Best, R.B., et al, Can non-mechanical proteins withstand force? Stretching barnase by atomic force microscopy and molecular dynamics simulation. Biophysical Journal, 2001. 81(4): pp. 2344–2356.CrossRefADSGoogle Scholar
  64. 64.
    Toofanny, R.D., P.M. Williams, and R. Elber, Long time-scale simulations of protein unfolding under force using the stochastic difference equation in length algorithm. Biophysical Journal, 2005. 88(1): pp. 185A-185A.Google Scholar
  65. 65.
    Zhang, X.H., D.F. Bogorin, and V.T. Moy, Molecular basis of the dynamic strength of the sialyl Lewis X-selectin interaction. Chemphyschem, 2004. 5(2): pp. 175–182.CrossRefGoogle Scholar
  66. 66.
    Zhang, X.H., et al, Molecular basis for the dynamic strength of the integrin alpha(4)beta(1)/VCAM-1 interaction. Biophysical Journal, 2004. 87(5): pp. 3470–3478.CrossRefADSGoogle Scholar
  67. 67.
    Bayas, M.V., et al, Lifetime measurements reveal kinetic differences between homophilic cadherin bonds. Biophysical Journal, 2006. 90(4): pp. 1385–1395.CrossRefADSGoogle Scholar
  68. 68.
    Toofanny, R.D. and P.M. Williams, Simulations of multi-directional forced unfolding of titin I27. Journal of Molecular Graphics & Modelling, 2006. 24(5): pp. 396–403.CrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media, LLC 2008

Authors and Affiliations

  • Phil Williams
    • 1
  1. 1.Laboratory of Biophysics and Surface Analysis, School of PharmacyUniversity of Nottingham, University ParkNottinghamUK

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