Nanomechanics of Yeast Surfaces Revealed by AFM

  • Etienne Dague
  • Audrey Beaussart
  • David Alsteens
Part of the NanoScience and Technology book series (NANO)


Despite the large and well-documented characterization of the microbial cell wall in terms of chemical composition, the determination of the mechanical properties of surface molecules in relation to their function remains a key challenge in cell biology.The emergence of powerful tools allowing molecular manipulations has already revolutionized our understanding of the surface properties of fungal cells. At the frontier between nanophysics and molecular biology, atomic force microscopy (AFM), and more specifically single-molecule force spectroscopy (SMFS), has strongly contributed to our current knowledge of the cell wall organization and nanomechanical properties. However, due to the complexity of the technique, measurements on live cells are still at their infancy.In this chapter, we describe the cell wall composition and recapitulate the principles of AFM as well as the main current methodologies used to perform AFM measurements on live cells, including sample immobilization and tip functionalization.The current status of the progress in probing nanomechanics of the yeast surface is illustrated through three recent breakthrough studies. Determination of the cell wall nanostructure and elasticity is presented through two examples: the mechanical response of mannoproteins from brewing yeasts and elasticity measurements on lacking polysaccharide mutant strains. Additionally, an elegant study on force-induced unfolding and clustering of adhesion proteins located at the cell surface is also presented.


Atomic Force Microscopy Yeast Cell Wall Force Spectroscopy Brewing Yeast Nanomechanical Property 
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.



This work was supported by the National Foundation for Scientific Research (FNRS) and the Université Catholique de Louvain. D.A. is a postdoctoral researcher of the FNRS. E.D. is a researcher of Centre National de la Recherche Scientifique (CNRS). The authors thank Childérick Severac for careful and critical reading of the chapter.


  1. 1.
    T.J. Beveridge, Ultrastructure, chemistry and function of the bacterial cell wall. Int. Rev. Cytol. 72, 229–317 (1981)Google Scholar
  2. 2.
    J.G.H. Wessels, Wall growth, protein excretion and morphogenesis in fungi. New Phytol. 123, 397–413 (1993)Google Scholar
  3. 3.
    L.J. García-Rodríguez, R. Valle, Á. Durán, C. Roncero, Cell integrity signaling activation in response to hyperosmotic shock in yeast. FEBS Lett. 579, 6186–6190 (2005)Google Scholar
  4. 4.
    D.E. Levin, Cell wall integrity signaling in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 69, 262–291 (2005)Google Scholar
  5. 5.
    F.M. Klis, P. Mol, K. Hellingwerf, S. Brul, Dynamics of cell wall structure in Saccharomyces cerevisiae. FEMS Microbiol. Rev. 26, 239–256 (2002)Google Scholar
  6. 6.
    R. Rodicio, J.J. Heinisch, Together we are strong – cell wall integrity sensors in yeasts. Yeast 27, 531–540 (2010)Google Scholar
  7. 7.
    A.M. Dranginis, J.M. Rauceo, J.E. Coronado, P.N.A. Lipke, Biochemical guide to yeast adhesins: glycoproteins for social and antisocial occasions. Microbiol. Mol. Biol. Rev. 71, 282–294 (2007)Google Scholar
  8. 8.
    R. Lewin, Microbial adhesion is a sticky problem. Science 224, 375–377 (1984)Google Scholar
  9. 9.
    E.L. Florin, V.T. Moy, H.E. Gaub, Adhesion forces between individual ligand receptor pairs. Science 264, 415–417 (1994)Google Scholar
  10. 10.
    H.C. Van der Mei, B. Van de Belt-Grotter, H.J. Busscher, Implications of microbial adhesion to hydrocarbons for evaluating cell surface hydrophobicity 2. adhesion mechanisms. Colloids Surf. B. Biointerfaces 5, 117–126 (1995)Google Scholar
  11. 11.
    P. Sundstrom, Adhesion in Candida spp. Cell. Microbiol. 4, 461–469 (2002)Google Scholar
  12. 12.
    J.C. Kapteyn, H. Van Den Ende, F.M. Klis, The contribution of cell wall proteins to the organization of the yeast cell wall. Biochim. Biophys. Acta-Gen. Subj. 1426, 373–383 (1999)Google Scholar
  13. 13.
    M. Osumi, The ultrastructure of yeast: cell wall structure and formation. Micron 29, 207–233 (1998)Google Scholar
  14. 14.
    J.E Coronado, S. Mneimneh, S.L. Epstein, W.G. Qiu, P.N. Lipke, Conserved processes and lineage-specific proteins in fungal cell wall evolution. Eukaryot. Cell 6, 2269–2277 (2007)Google Scholar
  15. 15.
    P.N. Lipke, R. Ovalle, Cell wall architecture in yeast: new structure and new challenges. J. Bacteriol. 180, 3735–3740 (1998)Google Scholar
  16. 16.
    G. Lesage, H. Bussey, Cell wall assembly in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 70, 317–343 (2006)Google Scholar
  17. 17.
    M.L. Richard, A. Plaine, Comprehensive analysis of glycosylphosphatidylinositol-anchored proteins in Candida albicans. Eukaryot. Cell 6, 119–133 (2007)Google Scholar
  18. 18.
    R.A. Daniel, J. Errington, Control of cell morphogenesis in bacteria: two distinct ways to make a rod-shaped cell. Cell 113, 767–776 (2003)Google Scholar
  19. 19.
    R.D. Turner et al. , Peptidoglycan architecture can specify division planes in Staphylococcus aureus. Nat. Commun. 1, 10.1038/ncomms1025 (2010)Google Scholar
  20. 20.
    S.W. Hell, Far-field optical nanoscopy. Science 316, 1153–1158 (2007)Google Scholar
  21. 21.
    Z. Gitai, New fluorescence microscopy methods for microbiology: sharper, faster, and quantitative. Curr. Opin. Microbiol. 12, 341–346 (2009)Google Scholar
  22. 22.
    V.R.F. Matias, T.J. Beveridge, Cryo-electron microscopy reveals native polymeric cell wall structure in Bacillus subtilis 168 and the existence of a periplasmic space. Mol. Microbiol. 56, 240–251 (2005)Google Scholar
  23. 23.
    E.A. Evans, D.A. Calderwood, Forces and bond dynamics in cell adhesion. Science 316, 1148–1153 (2007)Google Scholar
  24. 24.
    C. Bustamante, J.C. Macosko, G.J.L. Wuite, Grabbing the cat by the tail: manipulating molecules one by one. Nat. Rev. Mol. Cell Biol. 1, 130–136 (2000)Google Scholar
  25. 25.
    M. Sotomayor, K. Schulten, Single-molecule experiments in vitro and in silico. Science 316, 1144–1148 (2007)Google Scholar
  26. 26.
    D.J. Muller, Y.F. Dufrêne, Atomic force microscopy as a multifunctional molecular toolbox in nanobiotechnology. Nat. Nanotechnol. 3, 261–269 (2008)Google Scholar
  27. 27.
    P. Hinterdorfer, Y.F. Dufrêne, Detection and localization of single molecular recognition events using atomic force microscopy. Nat. Methods 3, 347–355 (2006)Google Scholar
  28. 28.
    Y.F. Dufrêne, Towards nanomicrobiology using atomic force microscopy. Nat. Rev. Microbiol. 6, 674–680 (2008)Google Scholar
  29. 29.
    K.C. Neuman, A. Nagy Single-molecule force spectroscopy: optical tweezers, magnetic tweezers and atomic force microscopy. Nat. Methods 5, 491–505 (2008)Google Scholar
  30. 30.
    D.J. Muller, J. Helenius, D. Alsteens, Y.F. Dufrêne, Force probing surfaces of living cells to molecular resolution. Nat. Chem. Biol. 5, 383–390 (2009)Google Scholar
  31. 31.
    G. Binnig, C.F. Quate, C. Gerber, Atomic force microscope. Phys. Rev. Lett. 56, 930–933 (1986)Google Scholar
  32. 32.
    S. Scheuring, Y.F. Dufrêne, Atomic force microscopy: probing the spatial organization, interactions and elasticity of microbial cell envelopes at molecular resolution. Mol. Microbiol. 75, 1327–1336 (2010)Google Scholar
  33. 33.
    K. El Kirat, S. Morandat, Y.F. Dufrêne, Nanoscale analysis of supported lipid bilayers using atomic force microscopy. Biochim. Biophys. Acta-Biomembr. 1798, 750–765 (2010)Google Scholar
  34. 34.
    S.Y. Liu, Y.F. Wang, Application of AFM in microbiology: a review. Scanning 32, 61–73 (2010)Google Scholar
  35. 35.
    L.S. Dorobantu, M.R. Gray, Application of atomic force microscopy in bacterial research. Scanning 32, 74–96 (2010)Google Scholar
  36. 36.
    D.J. Muller, M. Krieg, D. Alsteens, Y.F. Dufrêne, New frontiers in atomic force microscopy: analyzing interactions from single-molecules to cells. Curr. Opin. Biotechnol. 20, 4–13 (2009)Google Scholar
  37. 37.
    E. Lesniewska, P.E. Milhiet, M.C. Giocondi, C. Le Grimellec, Atomic force microscope imaging of cells and membranes. Methods Cell Biol. 68, 51–65 (2002)Google Scholar
  38. 38.
    F. Gaboriaud, Y.F. Dufrêne, Atomic force microscopy of microbial cells: application to nanomechanical properties, surface forces and molecular recognition forces. Colloids Surf. B. Biointerfaces 54, 10–19 (2007)Google Scholar
  39. 39.
    W.F. Heinz, J.H. Hoh, Spatially resolved force spectroscopy of biological surfaces using the atomic force microscope. Trends Biotechnol. 17, 143–150 (1999)Google Scholar
  40. 40.
    H.J. Busscher et al. Intermolecular forces and enthalpies in the adhesion of Streptococcus mutans and an antigen I/II-deficient mutant to laminin films. J. Bacteriol. 189, 2988–2995 (2007)Google Scholar
  41. 41.
    C. Roduit et al. , Elastic membrane heterogeneity of living cells revealed by stiff nanoscale membrane domains. Biophys. J. 94, 1521–1532 (2008)Google Scholar
  42. 42.
    J. Helenius, C.P. Heisenberg, H.E. Gaub, D.J. Muller, Single-cell force spectroscopy. J. Cell Sci. 121, 1785–1791 (2008)Google Scholar
  43. 43.
    C. Rankl et al. , Multiple receptors involved in human rhinovirus attachment to live cells. Proc. Natl. Acad. Sci. U. S. A. 105, 17778–17783 (2008)Google Scholar
  44. 44.
    M. Gad, A. Ikai, Method for immobilizing microbial cells on gel surface for dynamic AFM studies. Biophys. J. 69, 2226–2233 (1995)Google Scholar
  45. 45.
    E. Dague et al. , Assembly of live micro organisms on microstructured PDMS stamps by convective/capillary deposition for AFM bio-experiments. Nanotechnology 22, 395102 (2011)Google Scholar
  46. 46.
    R.D. Turner, N.H. Thomson, J. Kirkham, D. Devine, Improvement of the pore trapping method to immobilize vital coccoid bacteria for high-resolution AFM: a study of Staphylococcus aureus. J. Microsc.-Oxf. 238, 102–110 (2010)Google Scholar
  47. 47.
    C.D. Frisbie, L.F. Rozsnyai, A. Noy, M.S. Wrighton, C.M. Lieber, Functional-group imaging by chemical force microscopy. Science 265, 2071–2074 (1994)Google Scholar
  48. 48.
    D. Alsteens, E. Dague, P.G. Rouxhet, A.R. Baulard, Y.F. Dufrêne, Direct measurement of hydrophobic forces on cell surfaces using AFM. Langmuir 23, 11977–11979 (2007)Google Scholar
  49. 49.
    G.U. Lee, L.A. Chrisey, R.J. Colton, Direct measurement of the forces between complementary strands of DNA. Science 266, 771–773 (1994)Google Scholar
  50. 50.
    A. Touhami, B. Hoffmann, A. Vasella, F.A. Denis, Y.F. Dufrêne, Probing specific lectin-carbohydrate interactions using atomic force microscopy imaging and force measurements. Langmuir 19, 1745–1751 (2003)Google Scholar
  51. 51.
    A. Berquand et al. , Antigen binding forces of single antilysozyme Fv fragments explored by atomic force microscopy. Langmuir 21, 5517–5523 (2005)Google Scholar
  52. 52.
    V. Dupres et al., Nanoscale mapping and functional analysis of individual adhesins on living bacteria. Nat. Methods 2, 515–520 (2005)Google Scholar
  53. 53.
    C. Verbelen, H.J. Gruber, Y.F. Dufrêne, The NTA-His6 bond is strong enough for AFM single-molecular recognition studies. J. Mol. Recognit. 20, 490–494 (2007)Google Scholar
  54. 54.
    F. Kienberger et al., Recognition force spectroscopy studies of the NTA-His6 bond. Single Mol. 1, 59–65 (2000)Google Scholar
  55. 55.
    P. Hinterdorfer, W. Baumgartner, H.J. Gruber, K. Schilcher, & H. Schindler, Detection and localization of individual antibody-antigen recognition events by atomic force microscopy. Proc. Natl. Acad. Sci. U. S. A. 93, 3477–3481 (1996)Google Scholar
  56. 56.
    S. Allen et al. , Detection of antigen-antibody binding events with the atomic force microscope. Biochemistry 36, 7457–7463 (1997)Google Scholar
  57. 57.
    C.K. Riener et al. , Heterobifunctional crosslinkers for tethering single ligand molecules to scanning probes. Anal. Chim. Acta 497, 101–114 (2003)Google Scholar
  58. 58.
    A. Ebner et al. , A new, simple method for linking of antibodies to atomic force microscopy tips. Bioconj. Chem. 18, 1176–1184 (2007)Google Scholar
  59. 59.
    R. Ros et al., Antigen binding forces of individually addressed single-chain Fv antibody molecules. Proc. Natl. Acad. Sci. U. S. A. 95, 7402–7405 (1998)Google Scholar
  60. 60.
    F. Schwesinger et al., Unbinding forces of single antibody-antigen complexes correlate with their thermal dissociation rates. Proc. Natl. Acad. Sci. U. S. A. 97, 9972–9977 (2000)Google Scholar
  61. 61.
    W. Baumgartner, N. Golenhofen, N. Grundhofer, J. Wiegand, D. Drenckhahn, Ca2 +  dependency of N-cadherin function probed by laser tweezer and atomic force microscopy. J. Neurosci. 23, 11008–11014 (2003)Google Scholar
  62. 62.
    C. Stroh et al., Single-molecule recognition imaging-microscopy. Proc. Natl. Acad. Sci. U. S. A. 101, 12503–12507 (2004)Google Scholar
  63. 63.
    W. Baumgartner et al., Cadherin interaction probed by atomic force microscopy. Proc. Natl. Acad. Sci. U. S. A. 97, 4005–4010 (2000)Google Scholar
  64. 64.
    A. Razatos, Y.-L. Ong, M.M. Sharma, G. Georgiou, Molecular determinants of bacterial adhesion monitored by atomic force microscopy. Proc. Natl. Acad. Sci. U. S. A. 95, 11059–11064 (1998)Google Scholar
  65. 65.
    Y.L. Ong, A. Razatos, G. Georgiou, M.M. Sharma, Adhesion forces between E-coli bacteria and biomaterial surfaces. Langmuir 15, 2719–2725 (1999)Google Scholar
  66. 66.
    W.R. Bowen, N. Hilal, R.W. Lovitt, C.J. Wright, Direct measurement of the force of adhesion of a single biological cell using an atomic force microscope. Colloids Surf. Physicochem. Eng. Aspects 136, 231–234 (1998)Google Scholar
  67. 67.
    S.K. Lower, M.F. Hochella, T.J. Beveridge, Bacterial recognition of mineral surfaces: nanoscale interactions between Shewanella and α-FeOOH. Science 292, 1360–1363 (2001)Google Scholar
  68. 68.
    M. Benoit, D. Gabriel, G. Gerisch, H.E. Gaub, Discrete interactions in cell adhesion measured by single-molecule force spectroscopy. Nat. Cell Biol. 2, 313–317 (2000)Google Scholar
  69. 69.
    A. Touhami, B. Nysten, Y.F. Dufrêne, Nanoscale mapping of the elasticity of microbial cells by atomic force microscopy. Langmuir 19, 4539 (2003)Google Scholar
  70. 70.
    D. Alsteens et al., Structure, cell wall elasticity and polysaccharide properties of living yeast cells, as probed by AFM. Nanotechnology 19, 384005 (2008)Google Scholar
  71. 71.
    P.B. Dengis, L.R. Nelissen, P.G. Rouxhet, Mechanisms of yeast flocculation: comparison of top- and bottom-fermenting strains Appl. Environ. Microbiol. 61, 718–728 (1995)Google Scholar
  72. 72.
    E. Dague et al., An atomic force microscopy analysis of yeast mutants defective in cell wall architecture. Yeast 27, 673–684 (2010)Google Scholar
  73. 73.
    E. Dague, R. Bittar, F. Durand, H. Martin-Hyken, J.M. François, An Atomic Force Microscopy analysis of yeast mutants defective in cell wall architecture. Yeast 27, 673–784 (2010)Google Scholar
  74. 74.
    R.J. Karreman et al., The stress response protein Hsp12p increases the flexibility of the yeast Saccharomyces cerevisiae cell wall. Biochim. Biophys. Acta (BBA) Proteins Proteomics 1774, 131–137 (2007)Google Scholar
  75. 75.
    A.E.X. Brown, D.E. Discher, Conformational changes and signaling in cell and matrix physics. Curr. Biol. 19, R781-R789 (2009)Google Scholar
  76. 76.
    V. Vogel, M. Sheetz, Local force and geometry sensing regulate cell functions. Nat. Rev. Mol. Cell Biol. 7, 265–275 (2006)Google Scholar
  77. 77.
    J.C. Friedland, M.H. Lee, D. Boettiger, Mechanically activated integrin switch controls alpha(5)beta(1) function. Science 323 642–644 (2009)Google Scholar
  78. 78.
    B. Geiger, J.P. Spatz, A.D. Bershadsky, Environmental sensing through focal adhesions. Nat. Rev. Mol. Cell Biol. 10, 21–33 (2009)Google Scholar
  79. 79.
    A.D. Bershadsky, M. Kozlov, B. Geiger Adhesion-mediated mechanosensitivity: a time to experiment, and a time to theorize. Curr. Opin. Cell Biol. 18, 472–481 (2006)Google Scholar
  80. 80.
    A.S. Smith, K. Sengupta, S. Goennenwein, U. Seifert, E. Sackmann, Force-induced growth of adhesion domains is controlled by receptor mobility. Proc. Natl. Acad. Sci. U. S. A. 105, 6906–6911 (2008)Google Scholar
  81. 81.
    M. Gonzalez, P.W.J. de Groot, F.M. Klis, P.N. Lipke, Glycoconjugate structure and function in fungal cell walls, in Microbial Glycobiology, ed. by A.P. Moran (Academic, San Diego, 2009), pp 169–183Google Scholar
  82. 82.
    L.L. Hoyer, The ALS gene family of Candida albicans. Trends Microbiol. 9, 176–180 (2001)Google Scholar
  83. 83.
    L.L. Hoyer, C.B. Green, S.H. Oh, X.M. Zhao, Discovering the secrets of the Candida albicans agglutinin-like sequence (ALS) gene family – a sticky pursuit. Med. Mycol. 46, 1–15 (2008)Google Scholar
  84. 84.
    X.M. Zhao et al., ALS3 and ALS8 represent a single locus that encodes a Candida albicans adhesin; functional comparisons between Als3p and Als1p. Microbiology-(UK) 150, 2415–2428 (2004)Google Scholar
  85. 85.
    C.J. Nobile et al., Critical role of Bcr1-dependent adhesins in C. albicans biofilm formation in vitro and in vivo. PLoS Path. 2, 636–649 (2006)Google Scholar
  86. 86.
    S.A. Klotz et al., Degenerate peptide recognition by Candida albicans adhesins Als5p and Als1p. Infect. Immun. 72, 2029–2034 (2004)Google Scholar
  87. 87.
    D.C. Sheppard et al., Functional and structural diversity in the Als protein family of Candida albicans. J. Biol. Chem. 279, 30480–30489 (2004)Google Scholar
  88. 88.
    J.M. Rauceo et al., Threonine-rich repeats increase fibronectin binding in the Candida albicans adhesin Als5p. Eukaryot. Cell 5, 1664–1673 (2006)Google Scholar
  89. 89.
    J.M. Rauceo et al., Global cell surface conformational shift mediated by a Candida albicans adhesin. Infect. Immun. 72, 4948–4955 (2004)Google Scholar
  90. 90.
    C.B. Ramsook et al., Yeast cell adhesion molecules have functional amyloid-forming sequences. Eukaryot. Cell 9, 393–404 (2010)Google Scholar
  91. 91.
    H.N. Otoo, K.G. Lee, W.G. Qiu, P.N. Lipke, Candida albicans Als adhesins have conserved amyloid-forming sequences. Eukaryot. Cell 7 768–782 (2008)Google Scholar
  92. 92.
    A.T. Frank et al., Structure and function of glycosylated tandem repeats from Candida albicans Als adhesins. Eukaryot. Cell 9, 405–414 (2010)Google Scholar
  93. 93.
    D. Alsteens et al., Unfolding Individual Als5p Adhesion Proteins on Live Cells. ACS Nano 3, 1677–1682 (2009)Google Scholar
  94. 94.
    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)Google Scholar
  95. 95.
    D. Alsteens, M.C. Garcia, P.N. Lipke, Y.F. Dufrene, Force-induced formation and propagation of adhesion nanodomains in living fungal cells. Proc. Natl. Acad. Sci. U. S. A. 107, 20744–20749 (2010)Google Scholar
  96. 96.
    A.F. Oberhauser, P.E. Marszalek, H.P. Erickson, J.M. Fernandez, The molecular elasticity of the extracellular matrix protein tenascin. Nature 393, 181–185 (1998)Google Scholar
  97. 97.
    P.E. Marszalek, A.F. Oberhauser, Y.P. Pang, J.M. Fernandez, Polysaccharide elasticity governed by chair-boat transitions of the glucopyranose ring. Nature 396, 661–664 (1998)Google Scholar
  98. 98.
    F. Oesterhelt et al., Unfolding pathways of individual bacteriorhodopsins. Science 288, 143–146 (2000)Google Scholar
  99. 99.
    G. Lee et al., Nanospring behaviour of ankyrin repeats. Nature 440, 246–249 (2006)Google Scholar
  100. 100.
    J.K.H. Horber, M.J. Miles, Scanning probe evolution in biology. Science 302, 1002–1005 (2003)Google Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2012

Authors and Affiliations

  • Etienne Dague
    • 1
  • Audrey Beaussart
    • 2
  • David Alsteens
    • 2
  1. 1.CNRS UPR8001, LAASToulouseFrance
  2. 2.Institute of Condensed Matter and NanoscienceUniversité Catholique de LouvainLouvain-la-NeuveBelgium

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