Advertisement

Bridging the Gap Between Single-Molecule Unbinding Properties and Macromolecular Rheology

  • Makoto TakemasaEmail author
  • Andrew N. Round
  • Marit Sletmoen
  • Bjørn Torger Stokke
Chapter
Part of the Soft and Biological Matter book series (SOBIMA)

Abstract

Conformation and interactions between biological macromolecules are crucial for the mechanical properties of biological soft matter. In this chapter, the method and applications of the mechanical characteristics at the single-molecule level, from a fundamental point of view, are described as basis for understanding aspects of rheology. Atomic force microscope (AFM) and optical tweezers can be applied to investigate mechanical properties and interactions of molecules in the single molecular level. The force between two molecules as a result of specific and/or non-specific interactions can be determined as a function of distance between two molecules. Selected examples for interactions in macromolecules were highlighted based on observations by AFM-based force spectroscopy. This includes polysaccharide pairs such as interactions among hydrophobically modified hydroxyethyl cellulose (HMHEC), between protein polysaccharides and mucin–alginate. The mechanism of physically cross-linked hydrogel formation, HMHEC–amylose gel and alginate gels was also discussed based on single molecular pair interactions. For slower bond formation systems, which may not be capable with normal dynamic force spectroscopy, slide contact force spectroscopy can be applied. For slower dissociation rate, Dudko–Hummer–Szabo model and Friddle–Noy–De Yoreo model can be used for the analysis as an extension of the Bell–Evans model. The relation between characteristic timescale of interaction estimated in the single molecular study and relaxation spectra in the mechanical properties obtained at the macroscopic scale is presented as a possible way forward in understanding the gap between the mechanical properties in macroscopic and microscopic scale.

Keywords

Rheology Molecular interactions Dynamic force spectroscopy Atomic force microscope (AFM) 

References

  1. 1.
    Gennes, P., Scaling Concepts in Polymer Physics. 1979: Cornell University PressGoogle Scholar
  2. 2.
    Eaton, P. and P. West, Atomic Force Microscopy. 2010: Oxford University PressGoogle Scholar
  3. 3.
    Janshoff, A., et al., Force spectroscopy of molecular systems – Single molecule spectroscopy of polymers and biomolecules. Angewandte Chemie-International Edition, 2000. 39(18): p. 3213–3237Google Scholar
  4. 4.
    Sarid, D. and V. Elings, Review of Scanning Force Microscopy. Journal of Vacuum Science & Technology B, 1991. 9(2): p. 431–437CrossRefGoogle Scholar
  5. 5.
    Giannotti, M.I. and G.J. Vancso, Interrogation of single synthetic polymer chains and polysaccharides by AFM-based force spectroscopy. Chemphyschem, 2007. 8(16): p. 2290–2307CrossRefGoogle Scholar
  6. 6.
    Ashkin, A., Acceleration and Trapping of Particles by Radiation Pressure. Physical Review Letters, 1970. 24(4): p. 156–159Google Scholar
  7. 7.
    Ashkin, A., et al., Observation of a Single-Beam Gradient Force Optical Trap for Dielectric Particles. Optics Letters, 1986. 11(5): p. 288–290CrossRefGoogle Scholar
  8. 8.
    Moffitt, J.R., et al., Recent advances in optical tweezers. Annual Review of Biochemistry, 2008. 77: p. 205–228CrossRefGoogle Scholar
  9. 9.
    Williams, M.C., Optical Tweezers: Measuring Piconewton Forces, in Single Molecule Techniques, P. Schwille, Editor. 2002, Biophysical SocietyGoogle Scholar
  10. 10.
    Grandbois, M., et al., How strong is a covalent bond? Science, 1999. 283(5408): p. 1727–1730.CrossRefGoogle Scholar
  11. 11.
    Lee, G.U., L.A. Chrisey, and R.J. Colton, Direct Measurement of the Forces between Complementary Strands of DNA. Science, 1994. 266(5186): p. 771–773.CrossRefGoogle Scholar
  12. 12.
    Ortiz, C. and G. Hadziioannou, Entropic elasticity of single polymer chains of poly(methacrylic acid) measured by atomic force microscopy. Macromolecules, 1999. 32(3): p. 780–787CrossRefGoogle Scholar
  13. 13.
    Marszalek, P.E., et al., Polysaccharide elasticity governed by chair-boat transitions of the glucopyranose ring. Nature, 1998. 396(6712): p. 661–664.CrossRefGoogle Scholar
  14. 14.
    Noy, A. and R.W. Friddle, Practical single molecule force spectroscopy: How to determine fundamental thermodynamic parameters of intermolecular bonds with an atomic force microscope. Methods, 2013. 60(2): p. 142–150.CrossRefGoogle Scholar
  15. 15.
    Kienberger, F., et al., Molecular recognition imaging and force spectroscopy of single biomolecules. Accounts of Chemical Research, 2006. 39(1): p. 29–36.CrossRefGoogle Scholar
  16. 16.
    Rackham, B.D., et al., Non-covalent duplex to duplex crosslinking of DNA in solution revealed by single molecule force spectroscopy. Organic & Biomolecular Chemistry, 2013. 11(48): p. 8340–8347CrossRefGoogle Scholar
  17. 17.
    Morris, S., S. Hanna, and M.J. Miles, The self-assembly of plant cell wall components by single-molecule force spectroscopy and Monte Carlo modelling. Nanotechnology, 2004. 15(9): p. 1296–1301CrossRefGoogle Scholar
  18. 18.
    Hugel, T., et al., Elasticity of single polyelectrolyte chains and their desorption from solid supports studied by AFM based single molecule force spectroscopy. Macromolecules, 2001. 34: p. 1039–1047CrossRefGoogle Scholar
  19. 19.
    Friedsam, C., H.E. Gaub, and R.R. Netz, Probing surfaces with single-polymer atomic force microscope experiments. Biointerphases, 2006. 1(1): p. MR1-MR21CrossRefGoogle Scholar
  20. 20.
    Takemasa, M., M. Sletmoen, and B.T. Stokke, Single Molecular Pair Interactions between Hydrophobically Modified Hydroxyethyl Cellulose and Amylose Determined by Dynamic Force Spectroscopy. Langmuir, 2009. 25(17): p. 10174–10182.CrossRefGoogle Scholar
  21. 21.
    Egermayer, M., M. Karlberg, and L. Piculell, Gels of hydrophobically modified ethyl (hydroxyethyl) cellulose cross-linked by amylose: Effects of hydrophobe architecture. Langmuir, 2004. 20(6): p. 2208–2214.CrossRefGoogle Scholar
  22. 22.
    Chronakis, I.S., M. Egermayer, and L. Piculell, Thermoreversible gels of hydrophobically modified hydroxyethyl cellulose cross-linked by amylose. Macromolecules, 2002. 35(10): p. 4113–4122CrossRefGoogle Scholar
  23. 23.
    Bell, G.I., Models for Specific Adhesion of Cells to Cells. Science, 1978. 200(4342): p. 618–627.CrossRefGoogle Scholar
  24. 24.
    Evans, E. and K. Ritchie, Strength of a weak bond connecting flexible polymer chains. Biophysical Journal, 1999. 76(5): p. 2439–2447CrossRefGoogle Scholar
  25. 25.
    Friedsam, C., M. Seitz, and H.E. Gaub, Investigation of polyelectrolyte desorption by single molecule force spectroscopy. Journal of Physics-Condensed Matter, 2004. 16(26): p. S2369-S2382CrossRefGoogle Scholar
  26. 26.
    Evans, E. and K. Ritchie, Dynamic strength of molecular adhesion bonds. Biophysical Journal, 1997. 72(4): p. 1541–1555CrossRefGoogle Scholar
  27. 27.
    Karlson, L., K. Thuresson, and B. Lindman, A rheological investigation of the complex formation between hydrophobically modified ethyl (hydroxy ethyl) cellulose and cyclodextrin. Carbohydrate Polymers, 2002. 50(3): p. 219–226.CrossRefGoogle Scholar
  28. 28.
    Hane, F.T., S.J. Attwood, and Z. Leonenko, Comparison of three competing dynamic force spectroscopy models to study binding forces of amyloid-beta (1–42). Soft Matter, 2014. 10(12): p. 1924–1930CrossRefGoogle Scholar
  29. 29.
    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): 108101.Google Scholar
  30. 30.
    Friddle, R.W., A. Noy, and J.J. De Yoreo, Interpreting the widespread nonlinear force spectra of intermolecular bonds. Proc. Natl. Acad. Sci. U. S. A., 2012. 109(34): p. 13573–13578.Google Scholar
  31. 31.
    Harada, A. and M. Kamachi, Complex-Formation between Poly(Ethylene Glycol) and Alpha-Cyclodextrin. Macromolecules, 1990. 23(10): p. 2821–2823.CrossRefGoogle Scholar
  32. 32.
    Wanunu, M., Nanopores: A journey towards DNA sequencing. Physics of Life Reviews, 2012. 9(2): p. 125–158CrossRefGoogle Scholar
  33. 33.
    Laszlo, A.H., et al., Decoding long nanopore sequencing reads of natural DNA. Nature Biotechnology, 2014. 32(8): p. 829–833.CrossRefGoogle Scholar
  34. 34.
    Baaken, G., et al., High-Resolution Size-Discrimination of Single Nonionic Synthetic Polymers with a Highly Charged Biological Nanopore. Acs Nano, 2015. 9(6): p. 6443–6449CrossRefGoogle Scholar
  35. 35.
    Wenz, G., B.H. Han, and A. Muller, Cyclodextrin rotaxanes and polyrotaxanes. Chemical Reviews, 2006. 106(3): p. 782–817.CrossRefGoogle Scholar
  36. 36.
    Dunlop, A., et al., Mapping the positions of beads on a string: dethreading rotaxanes by molecular force spectroscopy. Nanotechnology, 2008. 19(34): 345706Google Scholar
  37. 37.
    Ashcroft, B.A., et al., An AFM/rotaxane molecular reading head for sequence-dependent DNA structures. Small, 2008. 4(9): p. 1468–1475CrossRefGoogle Scholar
  38. 38.
    Shigekawa, H., et al., The molecular abacus: STM manipulation of cyclodextrin necklace. Journal of the American Chemical Society, 2000. 122(22): p. 5411–5412CrossRefGoogle Scholar
  39. 39.
    Brough, B., et al., Evaluation of synthetic linear motor-molecule actuation energetics. Proceedings of the National Academy of Sciences of the United States of America, 2006. 103(23): p. 8583–8588CrossRefGoogle Scholar
  40. 40.
    Lussis, P., et al., A single synthetic small molecule that generates force against a load. Nature Nanotechnology, 2011. 6(9): p. 553–557CrossRefGoogle Scholar
  41. 41.
    Voulgarakis, N.K., et al., Sequencing DNA by dynamic force spectroscopy: Limitations and prospects. Nano Letters, 2006. 6(7): p. 1483–1486CrossRefGoogle Scholar
  42. 42.
    Qamar, S., P.M. Williams, and S.M. Lindsay, Can an atomic force microscope sequence DNA using a nanopore? Biophysical Journal, 2008. 94(4): p. 1233–1240.CrossRefGoogle Scholar
  43. 43.
    Larobina, D. and L. Cipelletti, Hierarchical cross-linking in physical alginate gels: a rheological and dynamic light scattering investigation. Soft Matter, 2013. 9(42): p. 10005–10015CrossRefGoogle Scholar
  44. 44.
    Siviello, C., F. Greco, and D. Larobina, Analysis of linear viscoelastic behaviour of alginate gels: effects of inner relaxation, water diffusion, and syneresis. Soft Matter, 2015. 11(30): p. 6045–6054.CrossRefGoogle Scholar
  45. 45.
    Mansel, B.W., et al., Zooming in: Structural Investigations of Rheologically Characterized Hydrogen-Bonded Low-Methoxyl Pectin Networks. Biomacromolecules, 2015. 16: p. 3209–3216.CrossRefGoogle Scholar
  46. 46.
    Stokke, B.T., et al., Distribution of Uronate Residues in Alginate Chains in Relation to Alginate Gelling Properties. Macromolecules, 1991. 24(16): p. 4637–4645CrossRefGoogle Scholar
  47. 47.
    Round, A.N., et al, in PreparationGoogle Scholar
  48. 48.
    S. Ballance et al., Preparation of high purity monodisperse oligosaccharides derived from mannuronan by size-exclusion chromatography followed by semi-preparative high-performance anion-exchange chromatography with pulsed amperometric detection. Carbohydrate Research 2009, 344(2), p.~255–259.Google Scholar
  49. 49.
    Rico, P., et al., High-Speed Force Spectroscopy Unfolds Titin at the Velocity of Molecular Dynamics Simulations. Science, 2013. 342(6159): p. 741–743.Google Scholar
  50. 50.
    Suresh, S.J. and V.M. Naik, Hydrogen bond thermodynamic properties of water from dielectric constant data. Journal of Chemical Physics, 2000. 113(21): p. 9727–9732.CrossRefGoogle Scholar
  51. 51.
    Hakem, I.F., et al., Temperature, pressure, and isotope effects on the structure and properties of liquid water: A lattice approach. Journal of Chemical Physics, 2007. 127(22): p.~224106.Google Scholar
  52. 52.
    Borgogna, M., et al., On the Initial Binding of Alginate by Calcium Ions. The Tilted Egg-Box Hypothesis. Journal of Physical Chemistry B, 2013. 117(24): p. 7277–7282CrossRefGoogle Scholar
  53. 53.
    Borukhov, I., et al., Elastically driven linker aggregation between two semiflexible polyelectrolytes. Physical Review Letters, 2001. 86(10): p. 2182–2185CrossRefGoogle Scholar
  54. 54.
    Corfield, A.P., Mucins: A biologically relevant glycan barrier in mucosal protection. Biochimica Et Biophysica Acta-General Subjects, 2015. 1850(1): p. 236–252CrossRefGoogle Scholar
  55. 55.
    Hang, H.C. and C.R. Bertozzi, The chemistry and biology of mucin-type O-linked glycosylation. Bioorganic & Medicinal Chemistry, 2005. 13(17): p. 5021–5034CrossRefGoogle Scholar
  56. 56.
    Liu, Z.H., et al., Polysaccharides-based nanoparticles as drug delivery systems. Advanced Drug Delivery Reviews, 2008. 60(15): p. 1650–1662CrossRefGoogle Scholar
  57. 57.
    Lai, S.K., Y.Y. Wang, and J. Hanes, Mucus-penetrating nanoparticles for drug and gene delivery to mucosal tissues. Advanced Drug Delivery Reviews, 2009. 61(2): p. 158–171CrossRefGoogle Scholar
  58. 58.
    Fuongfuchat, A., et al., Rheological studies of the interaction of mucins with alginate and polyacrylate. Carbohydrate Research, 1996. 284(1): p. 85–99CrossRefGoogle Scholar
  59. 59.
    Haugstad, K.E., et al., Direct Determination of Chitosan-Mucin Interactions Using a Single-Molecule Strategy: Comparison to Alginate-Mucin Interactions. Polymers, 2015. 7(2): p. 161–185CrossRefGoogle Scholar
  60. 60.
    Menchicchi, B., et al., Biophysical Analysis of the Molecular Interactions between Polysaccharides and Mucin. Biomacromolecules, 2015. 16(3): p. 924–935CrossRefGoogle Scholar
  61. 61.
    Taylor, C., et al., Rheological characterisation of mixed gels of mucin and alginate. Carbohydrate Polymers, 2005. 59(2): p. 189–195CrossRefGoogle Scholar
  62. 62.
    Nordgard, C.T. and K.I. Draget, Oligosaccharides As Modulators of Rheology in Complex Mucous Systems. Biomacromolecules, 2011. 12(8): p. 3084–3090.CrossRefGoogle Scholar
  63. 63.
    Sletmoen, M., et al., Oligoguluronate induced competitive displacement of mucin-alginate interactions: relevance for mucolytic function. Soft Matter, 2012. 8(32): p. 8413–8421CrossRefGoogle Scholar
  64. 64.
    Popeski-Dimovski, R., Work of adhesion between mucin macromolecule and calcium-alginate gels on molecular level. Carbohydrate Polymers, 2015. 123: p. 146–149CrossRefGoogle Scholar
  65. 65.
    Bucior, I. and M.M. Burger, Carbohydrate-carbohydrate interactions in cell recognition. Current Opinion in Structural Biology, 2004. 14(5): p. 631–637CrossRefGoogle Scholar
  66. 66.
    Haugstad, K.E., et al., Enhanced Self-Association of Mucins Possessing the T and Tn Carbohydrate Cancer Antigens at the Single-Molecule Level. Biomacromolecules, 2012. 13(5): p. 1400–1409.CrossRefGoogle Scholar
  67. 67.
    Haugstad, K.E., et al., Single molecule study of heterotypic interactions between mucins possessing the Tn cancer antigen. Glycobiology, 2015. 25(5): p. 524–534.CrossRefGoogle Scholar
  68. 68.
    Taylor, C., et al., The gel matrix of gastric mucus is maintained by a complex interplay of transient and nontransient associations. Biomacromolecules, 2003. 4(4): p. 922–927CrossRefGoogle Scholar
  69. 69.
    Spruijt, E., M.A.C. Stuart, and J. van der Gucht, Linear Viscoelasticity of Polyelectrolyte Complex Coacervates. Macromolecules, 2013. 46(4): p. 1633–1641CrossRefGoogle Scholar
  70. 70.
    Thünemann, A.F., et al., Polyelectrolyte complexes. Adv. Polym. Sci., 2004. 166: p. 113–171CrossRefGoogle Scholar
  71. 71.
    Bucur, C.B., Z. Sui, and J.B. Schlenoff, Ideal mixing in polyelectrolyte complexes and multilayers: Entropy driven assembly. Journal of the American Chemical Society, 2006. 128(42): p. 13690–13691CrossRefGoogle Scholar
  72. 72.
    Arents, G. and E.N. Moudrianakis, Topography of the histone octamer surface: repeating structural motifs utilized in the docking of nucleosomal DNA. Proceedings of the National Academy of Sciences, 1993. 90(22): p. 10489–10493CrossRefGoogle Scholar
  73. 73.
    Schiessel, H., The physics of chromatin. J. Phys. Condens. Matter, 2003. 15: p. R699–R774CrossRefGoogle Scholar
  74. 74.
    Mangenot, S., et al., Salt-induced conformation and interaction changes of nucleosome core particles. Biophysics Journal, 2002. 82: p. 345–356CrossRefGoogle Scholar
  75. 75.
    Mangenot, S., et al., Interactions between isolated nucleosome core particles. Eur. Phys. J. E, 2002. 7: p. 221–231Google Scholar
  76. 76.
    Bertin, A., et al., Role of histone tails in the conformation and interactions of nucleosome core particles. Biochemistry, 2004. 43: p. 4773–4780CrossRefGoogle Scholar
  77. 77.
    Müller, M., Polyelectrolyte Complexes in the Dispersed and Solid State II: Application Aspects. Advances in Polymer Science. Vol. 256. 2014, Berlin, Heidelberg: Springer Berlin Heidelberg. VII, 264 s. 137 illus., 1 illus. in color. : online resource.Google Scholar
  78. 78.
    Bertin, A., Polyelectrolyte complexes of DNA and polycations as gene delivery vectors. Advances in Polymer Science, 2014. 256: p. 103–196CrossRefGoogle Scholar
  79. 79.
    Müller, M., Sizing, shaping and pharmaceutical applications of polyelectrolyte complex nanoparticles. Advances in Polymer Science, 2014. 256: p. 197–260CrossRefGoogle Scholar
  80. 80.
    Hud, N.V. and K.H. Downing, Cryoelectron microscopy of l-phage DNA condensates in vitreous ice: The fine structure of DNA toroids. Proc. Natl. Acad. Sci. U. S. A., 2001. 98: p. 14925–14930CrossRefGoogle Scholar
  81. 81.
    Maurstad, G. and B.T. Stokke, Metastable and stable states of xanthan polyelectrolyte complexes studied by atomic force microscopy. Biopolymers, 2004. 74: p. 199–213.CrossRefGoogle Scholar
  82. 82.
    Priftis, D. and M. Tirrell, Phase behaviour and complex coacervation of aqueous polypeptide solutions. Soft Matter, 2012. 8(36): p. 9396–9405CrossRefGoogle Scholar
  83. 83.
    Spruijt, E., et al., Direct measurement of the strength of single ionic bonds between hydrated charges. ACS Nano, 2012. 6(6): p. 5297–5303CrossRefGoogle Scholar

Copyright information

© Springer Japan 2017

Authors and Affiliations

  • Makoto Takemasa
    • 1
    Email author
  • Andrew N. Round
    • 2
  • Marit Sletmoen
    • 3
  • Bjørn Torger Stokke
    • 4
  1. 1.School of Creative Science and EngineeringWaseda UniversityTokyoJapan
  2. 2.School of PharmacyUniversity of East Anglia, Norwich Research ParkNorwichUK
  3. 3.Department of BiotechnologyThe Norwegian University of Science and TechnologyTrondheimNorway
  4. 4.Department of PhysicsThe Norwegian University of Science and TechnologyTrondheimNorway

Personalised recommendations