Advertisement

Structural and Mechanical Mechanisms of Ocular Tissues Probed by AFM

  • Noël M. Ziebarth
  • Felix Rico
  • Vincent T. Moy
Chapter
Part of the NanoScience and Technology book series (NANO)

Summary

In recent years, the atomic force microscope (AFM) has become an important tool in ophthalmic research. It has gained popularity largely because AFM is not restricted by the diffraction limits of light microscopy and can be applied to resolve images with molecular resolution. AFM is a minimally invasive technique and can be used to visualize molecular structures under near-physiological conditions. In addition, the AFM can be employed as a force apparatus to characterize the viscoelastic properties of biomaterials on the micron level and at the level of individual proteins. In this article, we summarize recent AFM studies of ocular tissues, while highlighting the great potential of AFM technology in ophthalmic research. Previous research demonstrates the versatility of the AFM as high resolution imaging technique and as a sensitive force apparatus for probing the mechanical properties of ocular tissues. The structural and mechanical properties of ocular tissues are of major importance to the understanding of the optomechanical functions of the human eye. In addition, AFM has played an important role in the development and characterization of ocular biomaterials, such as contact lenses and intraocular lenses. Studying ocular tissues using Atomic Force Microscopy has enabled several advances in ophthalmic research.

Key words

Atomic force microscopy Nanoindentation Ophthalmology 

Preview

Unable to display preview. Download preview PDF.

Unable to display preview. Download preview PDF.

Notes

Acknowledgements

The authors thank Jean-Marie Parel, Ph.D., and Fabrice Manns, Ph.D., of the Ophthalmic Biophysics Center of Bascom Palmer Eye Institute, for providing their expertise in preparation of this chapter. The authors also thank the following support: NSF MRI 0722372; University of Miami SAC Award; NIH EY14225; NSF Graduate Student Fellowship (NMZ); Rakhi Jain, Ph.D., AMO, CA; Florida Lions Eye Bank; Vision Cooperative Research Centre, Sydney, New South Wales, Australia, supported by the Australian Federal Government through the Cooperative Research Centres Programme; NIH center grant P30-EY014801; Research to Prevent Blindness.

References

  1. 1.
    A. Engel, Y. Lyubchenko, and D. Muller, Atomic force microscopy: a powerful tool to observe biomolecules at work. Trends Cell Biol. 9(2), 77–80(1999).PubMedGoogle Scholar
  2. 2.
    W.F. Heinz, J.H. Hoh, Relative surface charge density mapping with the atomic force microscope. Biophys. J. 76(1 Pt 1), 528–538(1999).PubMedGoogle Scholar
  3. 3.
    H.W. Wu, T. Kuhn, and V.T. Moy, Mechanical properties of l929 cells measured by atomic force microscopy: effects of anticytoskeletal drugs and membrane crosslinking. Scanning 20, 389–397(1998).PubMedGoogle Scholar
  4. 4.
    A. Glasser, Restoration of accommodation. Curr. Opin. Ophthalmol. 17(1), 12–8(2006).MathSciNetGoogle Scholar
  5. 5.
    A. Glasser, Accommodation: mechanism and measurement. Ophthalmol. Clin. North Am. 19(1), 1–12(2006).PubMedGoogle Scholar
  6. 6.
    R.I. Barraquer, et al., Human lens capsule thickness as a function of age and location along the sagittal lens perimeter. Invest. Ophthalmol. Vis. Sci. 47(5), 2053–2060(2006).PubMedGoogle Scholar
  7. 7.
    D. Atchison, G. Smith (eds.), Optics of the Human Eye (Butterworth Heinemann: Oxford, UK, 2000), pp. 16–18.Google Scholar
  8. 8.
    A Glasser, P.L. Kaufman, Accommodation and presbyopia. In Adler’s Physiology of the Eye. Clinical Application, 10th edn., edited by P.L. Kaufman, A. Alm (Mosby, St Louis, MO, 2003), pp. 197–233.Google Scholar
  9. 9.
    R. Fisher, The significance of the shape of the lens and capsular energy changes in accommodation. J. Physiol. 201, 21–47(1969).PubMedGoogle Scholar
  10. 10.
    W.N. Charman, The eye in focus: accommodation and presbyopia. Clin. Exp. Optom. 91(3), 207–225(2008).PubMedGoogle Scholar
  11. 11.
    H.J. Wyatt, Some aspects of the mechanics of accommodation. Vis. Res. 28, 75–86(1988).PubMedGoogle Scholar
  12. 12.
    G. Binnig, C.F. Quate, C. Gerber, Atomic force microscope. Phys. Rev. Lett. 56, 930–933(1986).ADSGoogle Scholar
  13. 13.
    H.X. You, J.M. Lau, S. Zhang, L. Yu, Atomic force microscopy imaging of living cells: a preliminary study of the disruptive effect of the cantilever tip on cell morphology. Ultramicroscopy 82, 297–305(2000).PubMedGoogle Scholar
  14. 14.
    E. A-Hassan, W. Heinz, M.D. Antonik, N.P. D’Costa, S. Nageswaran, C.A. Schoenenberger, J.H. Hoh, Relative microelastic mapping of living cells by atomic force microscopy. Biophys. J. 74, 1564–1578(1998).PubMedGoogle Scholar
  15. 15.
    R. Afrin, A. Ikai, Force profiles of protein pulling with or without cytoskeletal links studied by AFM. Biochem. Biophys. Res. Commun. 348, 238–244(2006).PubMedGoogle Scholar
  16. 16.
    J. Alcaraz, L. Buscemi, M. Grabulosa, X. Trepat, B. Fabry, R. Farre, D. Navajas, Microrheology of human lung epithelial cells measured by atomic force microscopy. Biophys. J. 84, 2071–2079(2003).PubMedADSGoogle Scholar
  17. 17.
    M. Benoit, H.E. Gaub, Measuring cell adhesion forces with the atomic force microscope at the molecular level. Cells Tissues Organs 172, 174–189(2002).PubMedGoogle Scholar
  18. 18.
    H.J. Butt, B. Cappella, M. Kappl, Force measurements with the atomic force microscope: technique, interpretation and applications. Surf. Sci. Rep. 59, 1–152(2005).ADSGoogle Scholar
  19. 19.
    G.T. Charras, M.A. Horton, Single cell mechanotransduction and its modulation analyzed by atomic force microscope indentation. Biophys. J. 82, 2970–2981(2002).PubMedADSGoogle Scholar
  20. 20.
    S.L. Crick, F.C. Yin, Assessing micromechanical properties of cells with atomic force microscopy: importance of the contact point. Biomech. Model. Mechanobiol. 6, 199–210(2007).PubMedGoogle Scholar
  21. 21.
    J. Domke, S. Dannohl, W.J. Parak, O. Muller, W.K. Aicher, M. Radmacher, Substrate dependent differences in morphology and elasticity of living osteoblasts investigated by atomic force microscopy. Colloids Surf. B Biointerfaces 19, 367–379(2000).PubMedGoogle Scholar
  22. 22.
    V. Dupres, F.D. Menozzi, C. Locht, B.H. Clare, N.L. Abbott, S. Cuenot, C. Bompard, D. Raze, Y.F. Dufrene, Nanoscale mapping and functional analysis of individual adhesins on living bacteria. Nat Methods 2, 515–520(2005).PubMedGoogle Scholar
  23. 23.
    A.J. Engler, F. Rehfeldt, S. Sen, D.E. Discher, Microtissue elasticity: measurements by atomic force microscopy and its influence on cell differentiation. Methods Cell. Biol. 83, 521–545(2007).PubMedGoogle Scholar
  24. 24.
    E.L. Florin, M. Rief, H. Lehmann, M. Ludwig, C. Dornmair, V.T. Moy, H.E. Gaub, Sensing specific molecular-interactions with the atomic-force microscope. Biosens. Bioelectron. 10, 895–901(1995).Google Scholar
  25. 25.
    M. Grandbois, W. Dettmann, M. Benoit, H.E. Gaub, Affinity imaging of red blood cells using an atomic force microscope. J. Histochem. Cytochem. 48, 719–724(2000).PubMedGoogle Scholar
  26. 26.
    W. Haberle, J K.H. Horber, G. Binnig, Force microscopy on living cells. J. Vac. Sci. Technol. B 9, 1210–1213(1991).Google Scholar
  27. 27.
    A. Hategan, R. Law, S. Kahn, D.E. Discher, Adhesively-tensed cell membranes: lysis kinetics and atomic force microscopy probing. Biophys. J. 85, 2746–2759(2003).PubMedGoogle Scholar
  28. 28.
    E. Henderson, P.G. Haydon, D.S. Sakaguchi, Actin filament dynamics in living glial cells imaged by atomic force microscopy. Science 257, 1944–1946(1992).PubMedADSGoogle Scholar
  29. 29.
    P. Hinterdorfer, Y.F. Dufrene, Detection and localization of single molecular recognition events using atomic force microscopy. Nat. Methods 3, 347–355(2006).PubMedGoogle Scholar
  30. 30.
    J.H. Hoh, C.A. Schoenenberger, Surface morphology and mechanical properties of MDCK monolayers by atomic force microscopy. J. Cell Sci. 107, 1105–1114(1994).PubMedGoogle Scholar
  31. 31.
    K. Hyonchol, H. Arakawa, T. Osada, A. Ikai, Quantification of fibronectin and cell surface interactions by AFM. Colloid Surf. B-Biointerfaces 25, 33–43(2002).Google Scholar
  32. 32.
    H. Kim, H. Arakawa, T. Osada, A. Ikai, Quantification of cell adhesion force with AFM: distribution of vitronectin receptors on a living MC3T3-E1 cell. Ultramicroscopy 97, 359–363(2003).PubMedGoogle Scholar
  33. 33.
    E. Kokkoli, S.E. Ochsenhirt, M. Tirrell, Collective and single-molecule interactions of alpha(5)beta(1) integrins. Langmuir 20, 2397–2404(2004).PubMedGoogle Scholar
  34. 34.
    M. Krieg, Y. Arboleda-Estudillo, P.H. Puech, J. Kafer, F. Graner, D.J. Muller, C.P. Heisenberg, Tensile forces govern germ-layer organization in zebrafish. Nat. Cell Biol. 10, 429–436(2008).PubMedGoogle Scholar
  35. 35.
    R. Lal, B. Drake, D. Blumberg, D.R. Saner, P.K. Hansma, S.C. Feinstein, Imaging real-time neurite outgrowth and cytoskeletal reorganization with an atomic force microscope. Am. J. Physiol.-Cell Physiol. 269, C275–C285(1995).Google Scholar
  36. 36.
    I. Lee, R.E. Marchant, Force measurements on platelet surfaces with high spatial resolution under physiological conditions. Colloid Surf. B-Biointerfaces 19, 357–365(2000).Google Scholar
  37. 37.
    Q.S. Li, G.Y. Lee, C.N. Ong, C.T. Lim, AFM indentation study of breast cancer cells. Biochem. Biophys. Res. Commun. (2008).Google Scholar
  38. 38.
    T. Ludwig, R. Kirmse, K. Poole, U.S. Schwarz, Probing cellular microenvironments and tissue remodeling by atomic force microscopy. Pflugers Arch. 456, 29–49(2008).PubMedGoogle Scholar
  39. 39.
    R.E. Mahaffy, S. Park, E. Gerde, J. Kas, C.K. Shih, Quantitative analysis of the viscoelastic properties of thin regions of fibroblasts using atomic force microscopy. Biophys. J. 86, 1777–1793(2004).PubMedADSGoogle Scholar
  40. 40.
    R.E. Mahaffy, C.K. Shih, F.C. MacKintosh, J. Kas, Scanning probe-based frequency-dependent microrheology of polymer gels and biological cells. Phys. Rev. Lett. 85, 880–883(2000).PubMedADSGoogle Scholar
  41. 41.
    J.C. Martens, M. Radmacher, Softening of the actin cytoskeleton by inhibition of myosin II. Pflugers Arch. 456, 95–100(2008).PubMedGoogle Scholar
  42. 42.
    A.B. Mathur, A.M. Collinsworth, W.M. Reichert, W.E. Kraus, G.A. Truskey, Endothelial, cardiac muscle and skeletal muscle exhibit different viscous and elastic properties as determined by atomic force microscopy. J. Biomech. 34, 1545–1553(2001).PubMedGoogle Scholar
  43. 43.
    A.B. Mathur, G.A. Truskey, W.M. Reichert, Atomic force and total internal reflection fluorescence microscopy for the study of force transmission in endothelial cells. Biophys. J. 78, 1725–1735(2000).PubMedGoogle Scholar
  44. 44.
    V.T. Moy, E.L. Florin, H.E. Gaub, Adhesive forces between ligand and receptor measured by AFM. Biophys. J. 66, A340(1994).Google Scholar
  45. 45.
    P.H. Puech, A. Taubenberger, F. Ulrich, M. Krieg, D.J. Muller, C.P. Heisenberg, Measuring cell adhesion forces of primary gastrulating cells from zebrafish using atomic force microscopy. J. Cell Sci. 118, 4199–4206(2005).PubMedGoogle Scholar
  46. 46.
    C.A.J. Putman, K.O. Vanderwerf, B.G. Degrooth, N.F. Vanhulst, J. Greve, Viscoelasticity of living cells allows high-resolution imaging by tapping mode atomic-force microscopy. Biophys. J. 67, 1749–1753(1994).PubMedADSGoogle Scholar
  47. 47.
    C.A.J. Putman, K.O. Vanderwerf, B.G. Degrooth, N.F. Vanhulst, F.B. Segerink, J. Greve, Atomic force microscope with integrated optical microscope for biological applications. Rev. Sci. Instr. 63, 1914–1917(1992).ADSGoogle Scholar
  48. 48.
    M. Radmacher, Measuring the elastic properties of biological samples with the AFM. IEEE Eng. Med. Biol. Magn. 16, 47–57(1997).Google Scholar
  49. 49.
    M. Radmacher, Studying the mechanics of cellular processes by atomic force microscopy. Methods Cell Biol. 83, 347–372(2007).PubMedGoogle Scholar
  50. 50.
    M. Radmacher, R.W. Tillmann, M. Fritz, H.E. Gaub, From molecules to cells: imaging soft samples with the atomic force microscope. Science 257, 1900–1905(1992).PubMedADSGoogle Scholar
  51. 51.
    F. Rico, P. Roca-Cusachs, N. Gavara, R. Farre, M. Rotger, D. Navajas, Probing mechanical properties of living cells by atomic force microscopy with blunted pyramidal cantilever tips. Phys. Rev. E 72(2005).Google Scholar
  52. 52.
    F. Rico, P. Roca-Cusachs, R. Sunyer, R. Farre, D. Navajas, Cell dynamic adhesion and elastic properties probed with cylindrical atomic force microscopy cantilever tips. J. Mol. Recognit. 20, 459–466(2007).PubMedGoogle Scholar
  53. 53.
    M. Rief, M. Gautel, F. Oesterhelt, J.M. Fernandez, H.E. Gaub, Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 276, 1109–1112(1997).PubMedGoogle Scholar
  54. 54.
    C. Rotsch, F. Braet, E. Wisse, M. Radmacher, AFM imaging and elasticity measurements on living rat liver macrophages. Cell Biol. Int. 21, 685–696(1997).PubMedGoogle Scholar
  55. 55.
    C. Rotsch, M. Radmacher, Drug-Induced changes of cytoskeletal structure and mechanics in fibroblasts: an atomic force microscopy study. Biophys. J. 78, 520–535(2000).PubMedADSGoogle Scholar
  56. 56.
    S.W. Schneider, P. Pagel, C. Rotsch, T. Danker, H. Oberleithner, M. Radmacher, A. Schwab, Volume dynamics in migrating epithelial cells measured with atomic force microscopy. Pflugers Arch.-Eur. J. Physiol. 439, 297–303(2000).Google Scholar
  57. 57.
    S.G. Shroff, D.R. Saner, R. Lal, Dynamic micromechanical properties of cultured rat atrial myocytes measured by atomic-force microscopy. Am. J. Physiol.-Cell Physiol. 38, C286–C292(1995).Google Scholar
  58. 58.
    A. Simon, M.-C. Durrieu, Strategies and results of atomic force microscopy in the study of cellular adhesion. Micron 37, 13(2006).Google Scholar
  59. 59.
    L. Sirghi, J. Ponti, F. Broggi, F. Rossi, Probing elasticity and adhesion of live cells by atomic force microscopy indentation. Eur. Biophys. J. 37, 935–945(2008).PubMedGoogle Scholar
  60. 60.
    M. Stolz, R. Raiteri, A.U. Daniels, M.R. VanLandingham, W. Baschong, U. Aebi, Dynamic elastic modulus of porcine articular cartilage determined at two different levels of tissue organization by indentation-type atomic force microscopy. Biophys. J. 86, 3269–3283(2004).PubMedADSGoogle Scholar
  61. 61.
    A. Touhami, B. Nysten, Y.F. Dufrene, Nanoscale mapping of the elasticity of microbial cells by atomic force microscopy. Langmuir 19, 4539–4543(2003).Google Scholar
  62. 62.
    H.X. You, L. Yu, Atomic force microscopy imaging of living cells: progress, problems and prospects. Methods Cell Sci. 21, 1–17(1999).PubMedGoogle Scholar
  63. 63.
    A.E. Pelling, M.A. Horton, An historical perspective on cell mechanics. Pflugers Arch. 456, 3–12(2008).PubMedGoogle Scholar
  64. 64.
    P. Roca-Cusachs, I. Almendros, R. Sunyer, N. Gavara, R. Farre, D. Navajas, Rheology of passive and adhesion-activated neutrophils probed by atomic force microscopy. Biophys. J. 91, 3508–3518(2006).PubMedADSGoogle Scholar
  65. 65.
    G.A. Abrams, S.S. Schaus, S.L. Goodman, P.F. Nealey, C.J. Murphy. Nanoscale topography of the corneal epithelial basement membrane and Descemet’s membrane of the human. Cornea 19(1), 57–64(2000).PubMedGoogle Scholar
  66. 66.
    A. Antunes, F.V. Gozzo, M.I. Borella, M. Nakamura, A.M.V. Safatle, P.S.M. Barros, H.E. Toma. Atomic force imaging of ocular tissues: morphological study of healthy and cataract lenses. In Modern Research and Educational Topics in Microscopy (Formatex, 2007a).Google Scholar
  67. 67.
    A. Antunes, F.V. Gozzo, M. Nakamura, A.M. Safatle, S.L. Morelhão, H.E. Toma, P.S. Barros. Analysis of the healthy rabbit lens surface using MAC mode atomic force microscopy. Micron 38(3), 286–290(2007b).PubMedGoogle Scholar
  68. 68.
    N. Buzhynskyy, J.F. Girmens, W. Faigle, S. Scheuring, Human cataract lens membrane at subnanometer resolution. J. Mol. Biol. 374, 162–169(2007a).PubMedGoogle Scholar
  69. 69.
    N. Buzhynskyy, R.K. Hite, T. Walz, S. Scheuring. The supramolecular architecture of junctional microdomains in native lens membranes. EMBO Rep. 8, 51–55(2007b).PubMedGoogle Scholar
  70. 70.
    J. Candiello, M. Balasubramani, E.M. Schreiber, G.J. Cole, U. Mayer, W. Halfter, H. Lin. Biomechanical properties of native basement membranes. FEBS J. 274(11), 2897–908(2007).PubMedGoogle Scholar
  71. 71.
    N.J. Fullwood, A. Hammiche, H.M. Pollock, D.J. Hourston, M. Song. Atomic force microscopy of the cornea and sclera. Curr. Eye Res. 14, 529–535(1995).PubMedGoogle Scholar
  72. 72.
    M. Lombardo, M.P. De Santo, G. Lombardo, R. Barberi, S. Serrao. Atomic force microscopy analysis of normal and photoablated porcine corneas. J. Biomech. 39(14), 2719–2724(2006a).PubMedGoogle Scholar
  73. 73.
    S. Lydataki, E. Lesniewska, M.K. Tsilimbaris, C. Le Grimellec, L. Rochette, J.P. Goudonnet, I.J. Pallikaris. Observation of the posterior endothelial surface of the rabbit cornea using atomic force microscopy. Cornea 22(7), 651–664(2003).PubMedGoogle Scholar
  74. 74.
    S.B. Mallick, A. Ivanisevic. Study of the morphological and adhesion properties of collagen fibers in the Bruch’s membrane. J. Phys. Chem. B. 109(41):19052–19055(2005).PubMedGoogle Scholar
  75. 75.
    S.B. Mallick, S. Bhagwandin, A. Ivanisevic. Characterization of collagen fibers in Bruch’s membrane using chemical force microscopy. Anal. Bioanal. Chem. 386(3), 652–657(2006).PubMedGoogle Scholar
  76. 76.
    S. Mangenot, N. Buzhynskyy, J.F. Girmens, S. Scheuring. Malformation of junctional microdomains in cataract lens membranes from a type II diabetes patient. Eur. J. Physiol. (2008).Google Scholar
  77. 77.
    P. Matteini, F. Sbrana, B. Tiribilli, R. Pini. Atomic force microscopy and transmission electron microscopy analyses of low-temperature laser welding of the cornea. Lasers Med. Sci.(2008). DOI 10.1007/s10103–008–0617–4.Google Scholar
  78. 78.
    K.M. Meek, N.J. Fullwood. Corneal and scleral collagens – a microscopist’s perspective. Micron 32, 261–272(2001).PubMedGoogle Scholar
  79. 79.
    D. Meller, K. Peters, K. Meller. Human cornea and sclera studied by atomic force microscopy. Cell Tissue Res. 288(1), 111–118(1997).PubMedGoogle Scholar
  80. 80.
    A. Miyagawa, M. Kobayashi, Y. Fujita, M. Nakamura, K. Hirano, K. Kobayashi, Y. Miyake, Surface topology of collagen fibrils associated with proteoglycans in mouse cornea and sclera. Japan J. Ophthalmol. 44, 591–595(2000).Google Scholar
  81. 81.
    A. Miyagawa, M. Kobayashi, Y. Fujita, O. Hamdy, K. Hirano, M. Nakamura, Y. Miyake, Surface ultrastructure of collagen fibrils and their association with proteoglycans in human cornea and sclera by atomic force microscopy and energy-filtering transmission electron microscopy. Cornea 20(6), 651–656(2001).PubMedGoogle Scholar
  82. 82.
    A. Nógrádi, B. Hopp, K. Révész, G. Szabó, Z. Bor, L. Kolozsvari, Atomic force microscopic study of the human cornea following excimer laser keratectomy. Exp. Eye Res. 70, 363–368(2000).PubMedGoogle Scholar
  83. 83.
    S. Scheuring, N. Buzhynskyy, S. Jaroslawski, R.P. Goncalves, R.K. Hite, T. Walz, Structural models of the supramolecular organization of AQP0 and connexons in junctional microdomains. J. Struct. Biol. 160, 385–394(2007).PubMedGoogle Scholar
  84. 84.
    M.K. Tsilimbaris, E. Lesniewska, S. Lydataki, C. Le Grimellec, J.P. Goudonnet, I.G. Pallikaris, The use of atomic force microscopy for the observation of corneal epithelium surface. Invest. Ophthalmol. Vis. Sci. 41(3), 680–686(2000).PubMedGoogle Scholar
  85. 85.
    S. Yamamoto, J. Hitomi, M. Shigeno, S. Sawaguchi, H. Abe, T. Ushiki Atomic force microscopic studies of isolated collagen fibrils of the bovine cornea and sclera. Arch. Histol. Cytol. 60(4), 371–378(1997).PubMedGoogle Scholar
  86. 86.
    S. Yamamoto, H. Hashizume, J. Hitomi, M. Shigeno, S. Sawaguchi, H. Abe, T. Ushiki, The subfibrillar arrangement of corneal and scleral collagen fibrils as revealed by scanning electron and atomic force microscopy. Arch. Histol. Cytol. 63(2), 127–135(2000).PubMedGoogle Scholar
  87. 87.
    S. Yamamoto, J. Hitomi, S. Sawaguchi, H. Abe, M. Shigeno, T. Ushiki. Observation of human corneal and scleral collagen fibrils by atomic force microscopy. Japan J. Ophthalmol. 46, 496–501(2002).Google Scholar
  88. 88.
    N.M. Ziebarth, E.P. Wojcikiewicz, F. Manns, V.T. Moy, J.-M. Parel. Atomic force microscopy measurements of lens elasticity in monkey eyes. Mol. Vis. 13, 504–510(2007).PubMedGoogle Scholar
  89. 89.
    N.A. Burnham, O.P. Behrend, F. Oulevey, G. Gremaud, P.J. Gallo, D. Gourdon, E. Dupas, A.J. Kulik, H.M. Pollock, G.A.D. Briggs, How does a tip tap? Nanotechnology 8, 67–75(1997).ADSGoogle Scholar
  90. 90.
    J.H. Hoh, R. Lal, S.A. John, J.P. Revel, M.F. Arnsdorf, Atomic force microscopy and dissection of gap junctions. Science 253, 1405–1408(1991).PubMedADSGoogle Scholar
  91. 91.
    J.H. Hoh, G.E. Sosinsky, J.P. Revel, P.K. Hansma, Structure of the extracellular surface of the gap junction by atomic force microscopy. Biophys. J. 65, 149–163(1993).PubMedADSGoogle Scholar
  92. 92.
    P.K. Hansma, J.P. Cleveland, M. Radmacher, D.A. Walters, P.E. Hillner, M. Bezanilla, M. Fritz, D. Vie, H.G. Hansma, C.B. Prater, J. Massie, L. Fukunaga, J. Gurley, V. Elings, Tapping mode atomic force microscopy in liquids. Appl. Phys. Lett. 64, 1738–1740(1994).ADSGoogle Scholar
  93. 93.
    E. Nagao, J.A. Dvorak, An integrated approach to the study of living cells by atomic force microscopy. J. Microsc.-Oxf. 191, 8–19(1998).Google Scholar
  94. 94.
    M.A. Poggi, E.D. Gadsby, L.A. Bottomley, W.P. King, E. Oroudjev, H. Hansma, Scanning probe microscopy. Anal. Chem. 76, 3429–3443(2004).Google Scholar
  95. 95.
    B. Cappella, G. Dietler, Force-distance curves by atomic force microscopy. Surf. Sci. Rep. 34, 1(1999).Google Scholar
  96. 96.
    N.J. Tao, S.M. Lindsay, S. Lees, Measuring the microelastic properties of biological-material. Biophys. J. 63, 1165–1169(1992).PubMedADSGoogle Scholar
  97. 97.
    P. Frederix, T. Akiyama, U. Staufer, C. Gerber, D. Fotiadis, D.J. Muller, A. Engel, Atomic force bio-analytics. Curr. Opin. Chem. Biol. 7, 647(2003).Google Scholar
  98. 98.
    D.C. Lin, E.K. Dimitriadis, F. Horkay, Robust strategies for automated AFM force curve analysis-II: adhesion-influenced indentation of soft, elastic materials. J. Biomech. Eng. 129, 904–912(2007).PubMedGoogle Scholar
  99. 99.
    D.C. Lin, E.K. Dimitriadis, F. Horkay, Robust strategies for automated AFM force curve analysis–I. Non-adhesive indentation of soft, inhomogeneous materials. J. Biomech. Eng. 129, 430–440(2007).PubMedGoogle Scholar
  100. 100.
    F. Obataya, C. Nakamura, S.W. Han, N. Nakamura, J. Miyake, Mechanical sensing of the penetration of various nanoneedles into a living cell using atomic force microscopy. Biosens. Bioelectron. 20, 1652–1655(2005).PubMedGoogle Scholar
  101. 101.
    E.P. Wojcikiewicz, X. Zhang, V.T. Moy, Force and compliance measurements on living cells using atomic force microscopy (AFM). Biol. Proc. Online 6, 1–9(2004).Google Scholar
  102. 102.
    E.P. Wojcikiewicz, X. Zhang, A. Chen, V.T. Moy, Contributions of molecular binding events and cellular compliance to the modulation of leukocyte adhesion. J. Cell Sci. 116, 2531–2539(2003).PubMedGoogle Scholar
  103. 103.
    C.Y. Zhang, Y.W. Zhang, Computational analysis of adhesion force in the indentation of cells using atomic force microscopy. Phys. Rev. 77, 021912(2008).ADSGoogle Scholar
  104. 104.
    N.M. Ziebarth, E.P. Wojcikiewicz, F. Manns, V.T. Moy, J.M. Parel, Atomic force microscopy measurements of lens elasticity in monkey eyes. Mol. Vis. 13, 504–510(2007).PubMedGoogle Scholar
  105. 105.
    Y.F. Dufrene, P. Hinterdorfer, Recent progress in AFM molecular recognition studies. Pflugers Arch. 456, 237–245(2008).PubMedGoogle Scholar
  106. 106.
    V. Dupres, C. Verbelen, Y.F. Dufrene, Probing molecular recognition sites on biosurfaces using AFM. Biomaterials 28, 2393–2402(2007).PubMedGoogle Scholar
  107. 107.
    C. Verbelen, N. Christiaens, D. Alsteens, V. Dupres, A.R. Baulard, Y.F. Dufrene, Molecular mapping of lipoarabinomannans on mycobacteria. Langmuir (2009).Google Scholar
  108. 108.
    M. Gerhard, M.A. Nabil, Novel optical approach to atomic force microscopy. Appl. Phys. Lett. 53, 1045–1047(1988).Google Scholar
  109. 109.
    S. Alexander, L. Hellemans, O. Marti, J. Schneir, V. Elings, P. K. Hansma, M. Longmire, J. Gurley, An atomic-resolution atomic-force microscope implemented using an optical-lever. J. Appl. Phys. 65, 164–167(1989).ADSGoogle Scholar
  110. 110.
    M.A. Horton, G.T. Charras, G. Ballestrem, P.P. Lehenkari, Integration of atomic force and confocal microscopy. Single Mol. 1, 135–137(2000).ADSGoogle Scholar
  111. 111.
    J.L. Choy, S.H. Parekh, O. Chaudhuri, A.P. Liu, C. Bustamante, M.J. Footer, J.A. Theriot, D.A. Fletcher, Differential force microscope for long time-scale biophysical measurements. Rev. Sci. Instrum. 78, 043711(2007).PubMedADSGoogle Scholar
  112. 112.
    V. Heinrich, C. Ounkomol, Force versus axial deflection of pipette-aspirated closed membranes. Biophys. J. 93, 363–372(2007).PubMedADSGoogle Scholar
  113. 113.
    C. Ounkomol, H. Xie, P.A. Dayton, V. Heinrich, Versatile horizontal force probe for mechanical tests on pipette-held cells, particles, and membrane capsules. Biophys. J. 96, 1218–1231(2009).PubMedADSGoogle Scholar
  114. 114.
    P.P. Lehenkari, G.T. Charras, A. Nykanen, M.A. Horton, Adapting atomic force microscopy for cell biology. Ultramicroscopy 82, 289–295(2000).PubMedGoogle Scholar
  115. 115.
    P.H. Puech, K. Poole, D. Knebel, D.J. Muller, A new technical approach to quantify cell-cell adhesion forces by AFM. Ultramicroscopy (2006).Google Scholar
  116. 116.
    T. Ando, T. Uchihashi, N. Kodera, D. Yamamoto, A. Miyagi, M. Taniguchi, H. Yamashita, High-speed AFM and nano-visualization of biomolecular processes. Pflugers Arch. 456, 211–225(2008).PubMedGoogle Scholar
  117. 117.
    D.J. Muller, A. Engel, J.L. Carrascosa, M. Velez, The bacteriophage phi29 head-tail connector imaged at high resolution with the atomic force microscope in buffer solution. EMBO J. 16, 2547–2553(1997).PubMedGoogle Scholar
  118. 118.
    D.J. Muller, D. Fotiadis, S. Scheuring, S.A. Muller, A. Engel, Electrostatically balanced subnanometer imaging of biological specimens by atomic force microscope. Biophys. J. 76, 1101–1111(1999).PubMedGoogle Scholar
  119. 119.
    D.J. Muller, K.T. Sapra, S. Scheuring, A. Kedrov, P.L. Frederix, D. Fotiadis, A. Engel, Single-molecule studies of membrane proteins. Curr. Opin. Struct. Biol. 16, 489–495(2006).PubMedGoogle Scholar
  120. 120.
    G.T. Charras, M.A. Horton, Determination of cellular strains by combined atomic force microscopy and finite element modeling. Biophys. J. 83, 858–879(2002).PubMedADSGoogle Scholar
  121. 121.
    M.H. Abdulreda, V.T. Moy, Atomic force microscope studies of the fusion of floating lipid bilayers. Biophys. J. 92, 4369–4378(2007).PubMedADSGoogle Scholar
  122. 122.
    I. Obataya, C. Nakamura, S. Han, N. Nakamura, J. Miyake, Nanoscale operation of a living cell using an atomic force microscope with a nanoneedle. NanoLett. 5, 27–30(2005).ADSGoogle Scholar
  123. 123.
    L. Lu, S.J. Oswald, H. Ngu, F.C. Yin, Mechanical properties of actin stress fibers in living cells. Biophys. J. 95, 6060–6071(2008).PubMedADSGoogle Scholar
  124. 124.
    E.K. Dimitriadis, F. Horkay, J. Maresca, B. Kachar, R.S. Chadwick, Determination of elastic moduli of thin layers of soft material using the atomic force microscope. Biophys. J. 82, 2798–2810(2002).PubMedGoogle Scholar
  125. 125.
    K.D. Costa, A.J. Sim, F.C. Yin, Non-Hertzian approach to analyzing mechanical properties of endothelial cells probed by atomic force microscopy. J. Biomech. Eng. 128, 176–184(2006).PubMedGoogle Scholar
  126. 126.
    K.D. Costa, F.C.P. Yin, Analysis of indentation: implications for measuring mechanical properties with atomic force microscopy. J. Biomech. Eng. 121, 462–471(1999).PubMedGoogle Scholar
  127. 127.
    H. Hertz, On the contact of elastic bodies. In Hertz’s Miscellaneous Papers (Macmillan, London, 1881) pp. 146–162.Google Scholar
  128. 128.
    M. Radmacher, M. Fritz, C.M. Kacher, J.P. Cleveland, P.K. Hansma, Measuring the viscoelastic properties of human platelets with the atomic force microscope. Biophys. J. 70, 556–567(1996).PubMedGoogle Scholar
  129. 129.
    J.R. Barber, D.A. Billings, An approximate solution for the contact area and elastic compliance of a smooth punch of arbitrary shape. Int. J. Mech. Sci. 32, 991–997(1990).zbMATHGoogle Scholar
  130. 130.
    E.M. Darling, S. Zauscher, J.A. Block, F. Guilak, A thin-layer model for viscoelastic, stress-relaxation testing of cells using atomic force microscopy: do cell properties reflect metastatic potential? Biophys. J. 92, 1784–1791(2007).PubMedADSGoogle Scholar
  131. 131.
    B.A. Smith, B. Tolloczko, J.G. Martin, P. Grutter, Probing the viscoelastic behavior of cultured airway smooth muscle cells with atomic force microscopy: stiffening induced by contractile agonist. Biophys. J. 88, 2994–3007(2005).PubMedGoogle Scholar
  132. 132.
    H.F. Edelhauser, J.L. Ubels, Cornea and Sclera. In Adler’s Physiology of the Eye, edited by P. Kaufman, A. Alm (Mosby, Inc., St. Louis, MO, 2003).Google Scholar
  133. 133.
    F.E. Fantes, G.O. Waring, Effect of excimer laser radiant exposure on uniformity of ablated corneal surface. Lasers Surg. Med. 9(6), 533–542(1989).PubMedGoogle Scholar
  134. 134.
    T. Møller-Pedersen, H.D. Cavanagh, W.M. Petroll, J.V. Jester, Stromal wound healing explains refractive instability and haze development after photorefractive keratectomy. Ophthalmology. 107, 1235–1245(2000).PubMedGoogle Scholar
  135. 135.
    A. Keirl, C. Christie. Clinical Optics and Refraction: A Guide for Optometrists, Contact Lens Opticians and Dispensing Opticians. (Elsevier Health Sciences, New York, 2007).Google Scholar
  136. 136.
    E. Chalupa, H.A. Swarbrick, B.A. Holden, J. Sjöstrand, Severe corneal infections associated with contact lens wear. Ophthalmology. 94(1), 17–22(1987).PubMedGoogle Scholar
  137. 137.
    S. Bhatia, E.P. Goldberg, J.B. Enns, Examination of contact lens surfaces by atomic force microscope (AFM). CLAO J. 23(4), 264–269(1997).PubMedGoogle Scholar
  138. 138.
    J.M. González-Méijome, A. López-Alemany, J.B. Almeida, M.A. Parafita, M.F. Refojo, Microscopic observation of unworn siloxane–hydrogel soft contact lenses by atomic force microscopy. J. Biomed. Mater. Res. Part B: Appl. Biomater. 76B(2), 412–418 (2005).Google Scholar
  139. 139.
    J. Baguet, F. Sommer, T.M. Duc, Imaging surfaces of hydrophilic contact lenses with the atomic force microscope. Biomaterials. 14(4), 279–284(1993).PubMedGoogle Scholar
  140. 140.
    J. Baguet, F. Sommer, V. Claudon-Eyl, T.M. Duc, Characterization of lacrymal component accumulation on worn soft contact lens surfaces by atomic force microscopy. Biomaterials. 16(1), 3–9(1995).PubMedGoogle Scholar
  141. 141.
    E.P. Goldberg, S. Bhatia, J.B. Enns, Hydrogel contact lens-corneal interactions: a new mechanism for deposit formation and corneal injury. CLAO J. 23(4), 243–248(1997).PubMedGoogle Scholar
  142. 142.
    V. Guryca, R. Hobzová, M. Prádný, J. Sirc, J. Michálek. Surface morphology of contact lenses probed with microscopy techniques. Cont. Lens Anterior Eye. 30(4), 215–222(2007).PubMedGoogle Scholar
  143. 143.
    C. Maldonado-Codina, N. Efron, Impact of manufacturing technology and material composition on the surface characteristics of hydrogel contact lenses. Clin. Exp. Optom. 88(6), 396–404(2005).PubMedGoogle Scholar
  144. 144.
    C.E. Rabke, P.L. Valint, D.M. Ammon, Ophthalmic applications of atomic force microscopy. ICLC 22, 32–41(1995).Google Scholar
  145. 145.
    J.H. Teichroeb, J.A. Forrest, V. Ngai, J.W. Martin, L. Jones, J. Medley, Imaging protein deposits on contact lens materials. Optom. Vis. Sci. 85(12), 1151–1164(2008).Google Scholar
  146. 146.
    G.L. Grobe, P.L. Valint, D.M. Ammon, Surface chemical structure for soft contact lenses as a function of polymer processing. J. Biomed. Mater. Res. 32(1), 45–54(1996).PubMedGoogle Scholar
  147. 147.
    R.P. Santos, T.T. Arruda, C.B. Carvalho, V.A. Carneiro, L.Q. Braga, E.H. Teixeira, F.V. Arruda, B.S. Cavada, A. Havt, T.M. de Oliveira, G.A. Bezerra, V.N. Freire, Correlation between Enterococcus faecalis biofilms development stage and quantitative surface roughness using atomic force microscopy. Microsc. Microanal. 14(2), 150–158(2008).PubMedGoogle Scholar
  148. 148.
    G.M. Bruinsma, M. Rustema-Abbing, J. de Vries, H.J. Busscher, M.L. van der Linden, J.M. Hooymans, H.C. van der Mei, Multiple surface properties of worn RGP lenses and adhesion of Pseudomonas aeruginosa. Biomaterials 24(9), 1663–1670(2003).PubMedGoogle Scholar
  149. 149.
    A. Opdahl, S.H. Kim, T.S. Koffas, C. Marmo, G.A. Somorjai, Surface mechanical properties of pHEMA contact lenses: viscoelastic and adhesive property changes on exposure to controlled humidity. J. Biomed. Mater. Res. A. 67(1), 350–356(2003).PubMedGoogle Scholar
  150. 150.
    S.H. Kim, C. Marmo, G.A. Somorjai, Friction studies of hydrogel contact lenses using AFM: non-crosslinked polymers of low friction at the surface. Biomaterials 22(24), 3285–3294(2001).PubMedGoogle Scholar
  151. 151.
    S.H. Kim, A. Opdahl, C. Marmo, G.A. Somorjai, AFM and SFG studies of pHEMA-based hydrogel contact lens surfaces in saline solution: adhesion, friction, and the presence of non-crosslinked polymer chains at the surface. Biomaterials 23(7), 1657–1666(2002).PubMedGoogle Scholar
  152. 152.
    L.A. Remington, Clinical Anatomy of the Visual System (Butterworth-Heinemann, Boston, 1998).Google Scholar
  153. 153.
    R.C. Augusteyn, Growth of the lens: in vitro observations. Clin. Exp. Optometry 91(3), 226–239(2008).Google Scholar
  154. 154.
    S.K. Pandey, J. Thakur, L. Werner, M.E. Wilson, L.P. Werner, A.M. Izak, D.J. Apple, The human crystalline lens, ciliary body, and zonules: their relevance to presbyopia. In Presbyopia: A Surgical Textbook, edited by A. Agarwal (Slack Incorporated, Thorofare, NJ, 2002).Google Scholar
  155. 155.
    V.L. Taylor, K.J. Al-Ghoul, C.W. Lane, V.A. Davis, J.R. Kuszak, M.J. Costello, Morphology of the normal human lens. Invest. Ophthalmol. Visual Sci. 37(7), 1396–1410(1996).Google Scholar
  156. 156.
    D.C. Beebe, The lens. In Adler’s Physiology of the Eye, edited by P. Kaufman, A. Alm (Mosby, Inc., St. Louis, MO, 2003).Google Scholar
  157. 157.
    A. Shiels, S. Bassnett, Mutations in the founder of the MIP gene family underlie cataract development in the mouse. Nat. Genet. 12(2), 212–215(1996).PubMedGoogle Scholar
  158. 158.
    E.F. Fincham. The mechanism of accommodation. Br. J. Ophthalmol. Monograph Supplement VIII (1937).Google Scholar
  159. 159.
    J. Kessler, Experiments in refilling the lens. Arch. Ophthalmol. 71, 412–417(1964).PubMedGoogle Scholar
  160. 160.
    J. Kessler, Refilling the rabbit lens. Further experiments. Arch. Ophthalmol. 76(4), 596–598(1966).Google Scholar
  161. 161.
    R.F. Fisher, Elastic constants of the human lens capsule. J. Physiol. 201, 1–19(1969).PubMedGoogle Scholar
  162. 162.
    E.F. Fincham, The changes in the form of the crystalline lens in accommodation. Trans. Opt. Soc. 26, 239–269(1925).Google Scholar
  163. 163.
    B. Gilmartin, The aetiology of presbyopia: a summary of the role of lenticular and extralenticular structures. Opthal. Physiol. Opt. 15(5), 431–437(1995).Google Scholar
  164. 164.
    M. Tscherning. Le mecanisme de l’accommodation. Annals Oculiat. 131, 168–179(1904).Google Scholar
  165. 165.
    S. Krag, T. Olsen, T.T. Andreassen, Biomechanical characteristics of the human anterior lens capsule in relation to age. Invest. Ophthalmol. Visual Sci. 38, 357–363(1997).Google Scholar
  166. 166.
    K.R. Heys, S.L. Cram, R.J.W. Truscott, Massive increase in the stiffness of the human lens nucleus with age: the basis for presbyopia? Mol. Vis. 10, 956–963(2004).PubMedGoogle Scholar
  167. 167.
    K.R. Heys, R.J. Truscott, The stiffness of human cataract lenses is a function of both age and the type of cataract. Exp. Eye Res. 86(4), 701–703(2008).PubMedGoogle Scholar
  168. 168.
    H.A. Weeber, G. Eckert, F. Soergel, C.H. Meyer, W. Pechhold, R.G.L. van der Heijde. Dynamic mechanical properties of human lenses. Exp. Eye Res. 80, 425–434(2005).PubMedGoogle Scholar
  169. 169.
    F. Soergel, C. Meyer, G. Eckert, B. Abele, W. Pechhold, Spectral analysis of viscoelasticity of the human lens. J. Refract. Surg. 15, 714–716(1999).PubMedGoogle Scholar
  170. 170.
    H.A. Weeber, G. Eckert, W. Pechhold, R.G.L. van der Heijde, Stiffness gradient in the crystalline lens. Graefe’s Arch. Clin. Exp. Ophthalmol. 245, 1357–1366(2007).Google Scholar
  171. 171.
    R.A. Schachar, B.K. Pierscionek, Lens hardness not related to the age-related decline of accommodative amplitude. Mol. Vis. 13, 1010–1011(2007).PubMedGoogle Scholar
  172. 172.
    M. Lombardo, M.P. De Santo, G. Lombardo, R. Barberi, S. Serrao, Analysis of intraocular lens surface properties with atomic force microscopy. J. Cataract Refract. Surg. 32, 1378–1384(2006b).PubMedGoogle Scholar
  173. 173.
    Y. Ohnishi, T. Yoshitomi, T. Sakamoto, K. Fujisawa, T. Ishibashi. Evaluation of cellular adhesions on silicone and poly(methyl methacrylate) intraocular lenses in monkey eyes; an electron microscopic study. J. Cataract Refract. Surg. 27, 2036–2040(2001).PubMedGoogle Scholar
  174. 174.
    C. Cassinelli, M. Morra, A. Pavesio, D. Renier. Evaluation of interfacial properties of hyaluronan coated poly(methylmethacrylate) intraocular lenses. J. Biomater. Sci. Polym. Edn. 11(9), 961–977(2000).Google Scholar
  175. 175.
    D. Bozukova, C. Pagnoulle, M.C. De Pauw-Gillet, S. Desbief, R. Lazzaroni, N. Ruth, R. Jrme, C. Jrme. Improved performances of intraocular lenses by poly(ethylene glycol) chemical coatings. Biomacromolecules. 8(8), 2379–2387(2007).PubMedGoogle Scholar
  176. 176.
    M. Dogru, K. Tetsumoto, Y. Tagami, K. Kato, K. Nakamae. Optical and atomic force microscopy of an explanted AcrySof intraocular lens with glistenings. J. Cataract Refract. Surg. 26(4), 571–575(2000).PubMedGoogle Scholar
  177. 177.
    R.K. Sharma, B.E.J. Ehinger. Development and structure of the retina. In Adler’s Physiology of the Eye, edited by P. Kaufman, A. Alm. (Mosby, Inc., St. Louis, MO, 2003).Google Scholar
  178. 178.
    A. Hendrickson, Organization of the adult primate fovea. In Macular Degeneration, edited by P.L. Penfold, J.M. Provis (Springer, Berlin, 2005).Google Scholar
  179. 179.
    P.L. Penfold, J. Wong, D. van Driel, J.M. Provis, M.C. Madigan, Immunology and age-related macular degeneration. In Macular Degeneration, edited by P.L. Penfold, J.M. Provis (Springer, Berlin, 2005).Google Scholar
  180. 180.
    G. Wu, Retina: The Fundamentals (W.B. Saunders, St. Louis, MO, 1995).Google Scholar
  181. 181.
    W. Halfter, M. Willem, U. Mayer, Basement membrane-dependent survival of retinal ganglion cells. Invest. Ophthalmol. Visual Sci. 46, 1000–1009(2005).Google Scholar
  182. 182.
    R.F. Fisher, The elastic constants of the human lens. J. Physiol. 212, 147–180(1971).PubMedGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2010

Authors and Affiliations

  • Noël M. Ziebarth
    • 1
  • Felix Rico
    • 2
  • Vincent T. Moy
    • 3
  1. 1.Department of Biomedical EngineeringUniversity of Miami College of EngineeringCoral GablesUSA
  2. 2.Department of Physiology and BiophysicsUniversity of Miami Miller School of MedicineMiamiUSA
  3. 3.Department of Physiology and BiophysicsUniversity of MiamiMiamiUSA

Personalised recommendations