Atomic force microscopy-based characterization and design of biointerfaces

Abstract

Atomic force microscopy (AFM)-based methods have matured into a powerful nanoscopic platform, enabling the characterization of a wide range of biological and synthetic biointerfaces ranging from tissues, cells, membranes, proteins, nucleic acids and functional materials. Although the unprecedented signal-to-noise ratio of AFM enables the imaging of biological interfaces from the cellular to the molecular scale, AFM-based force spectroscopy allows their mechanical, chemical, conductive or electrostatic, and biological properties to be probed. The combination of AFM-based imaging and spectroscopy structurally maps these properties and allows their 3D manipulation with molecular precision. In this Review, we survey basic and advanced AFM-related approaches and evaluate their unique advantages and limitations in imaging, sensing, parameterizing and designing biointerfaces. It is anticipated that in the next decade these AFM-related techniques will have a profound influence on the way researchers view, characterize and construct biointerfaces, thereby helping to solve and address fundamental challenges that cannot be addressed with other techniques.

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Figure 1: AFM imaging principles and applications characterizing biointerfaces.
Figure 2: AFM-based force spectroscopy from single molecules to cells.
Figure 3: AFM-based imaging and mapping of mechanical properties of biointerfaces.
Figure 4: AFM-based imaging and affinity mapping of biointerfaces.
Figure 5: Characterizing reactions of biointerfaces in real time using AFM-based microsensors.
Figure 6: AFM-based sculpting, patterning and assembly of biointerfaces.
Figure 7: Combining AFM with other microscopic and spectroscopic approaches.

References

  1. 1

    Nel, A. E. et al. Understanding biophysicochemical interactions at the nano–bio interface. Nat. Mater. 8, 543–557 (2009).

  2. 2

    Ross, A. M. & Lahann, J. Current trends and challenges in biointerfaces science and engineering. Annu. Rev. Chem. Biomol. 6, 161–186 (2015).

  3. 3

    Stevens, M. M. & George, J. H. Exploring and engineering the cell surface interface. Science 310, 1135–1138 (2005).

  4. 4

    Andrews, R. N., Co, C. C. & Ho, C. C. Engineering dynamic biointerfaces. Curr. Opin. Chem. Eng. 11, 28–33 (2016).

  5. 5

    Gerber, C. & Lang, H. P. How the doors to the nanoworld were opened. Nat. Nanotechnol. 1, 3–5 (2006).

  6. 6

    Binnig, G., Quate, C. F. & Gerber, C. Atomic force microscope. Phys. Rev. Lett. 56, 930–933 (1986). This paper describes the invention of AFM.

  7. 7

    Drake, B. et al. Imaging crystals, polymers, and processes in water with the atomic force microscope. Science 243, 1586–1589 (1989).

  8. 8

    Radmacher, M., Tillmann, R. W., Fritz, M. & Gaub, H. E. From molecules to cells: imaging soft samples with the atomic force microscope. Science 257, 1900–1905 (1992).

  9. 9

    Garcia, R. & Herruzo, E. T. The emergence of multifrequency force microscopy. Nat. Nanotechnol. 7, 217–226 (2012).

  10. 10

    Ando, T., Uchihashi, T. & Kodera, N. High-speed AFM and applications to biomolecular systems. Ann. Rev. Biophys. 42, 393–414 (2013).

  11. 11

    Zhang, S., Aslan, H., Besenbacher, F. & Dong, M. Quantitative biomolecular imaging by dynamic nanomechanical mapping. Chem. Soc. Rev. 43, 7412–7429 (2014).

  12. 12

    Dufrêne, Y. F. et al. Imaging modes of atomic force microscopy for application in molecular and cell biology. Nat. Nanotechnol. http://dx.doi.org/10.1038/nnano.2017.45 (2017).

  13. 13

    Frisbie, C. D., Rozsnyai, L. F., Noy, A., Wrighton, M. S. & Lieber, C. M. Functional group imaging by chemical force microscopy. Science 265, 2071–2074 (1994).

  14. 14

    Hinterdorfer, P. & Dufrêne, Y. F. Detection and localization of single molecular recognition events using atomic force microscopy. Nat. Methods 3, 347–355 (2006).

  15. 15

    Müller, D. J., Helenius, J., Alsteens, D. & Dufrêne, Y. F. Force probing surfaces of living cells to molecular resolution. Nat. Chem. Biol. 5, 383–390 (2009).

  16. 16

    Dufrêne, Y. F., Martinez-Martin, D., Medalsy, I., Alsteens, D. & Müller, D. J. Multiparametric imaging of biological systems by force–distance curve-based AFM. Nat. Methods 10, 847–854 (2013).

  17. 17

    Lang, H. P. & Gerber, C. Microcantilever sensors. Top. Curr. Chem. 285, 1–27 (2008).

  18. 18

    Müller, D. J. & Dufrêne, Y. F. Atomic force microscopy as a multifunctional molecular toolbox in nanobiotechnology. Nat. Nanotechnol. 3, 261–269 (2008).

  19. 19

    Pires, D. et al. Nanoscale three-dimensional patterning of molecular resists by scanning probes. Science 328, 732–735 (2010).

  20. 20

    Garcia, R., Knoll, A. W. & Riedo, E. Advanced scanning probe lithography. Nat. Nanotechnol. 9, 577–587 (2014).

  21. 21

    Puchner, E. M. & Gaub, H. E. Single-molecule mechanoenzymatics. Ann. Rev. Biophys. 41, 497–518 (2012).

  22. 22

    Cattin, C. J. et al. Mechanical control of mitotic progression in single animal cells. Proc. Natl Acad. Sci. USA 112, 11258–11263 (2015).

  23. 23

    Kufer, S. K., Puchner, E. M., Gumpp, H., Liedl, T. & Gaub, H. E. Single-molecule cut-and-paste surface assembly. Science 319, 594–596 (2008).

  24. 24

    Engel, A. & Müller, D. J. Observing single biomolecules at work with the atomic force microscope. Nat. Struct. Biol. 7, 715–718 (2000).

  25. 25

    Kuznetsov, Y. G. & McPherson, A. Atomic force microscopy in imaging of viruses and virus-infected cells. Microbiol. Mol. Biol. Rev. 75, 268–285 (2011).

  26. 26

    Ido, S. et al. Beyond the helix pitch: direct visualization of native DNA in aqueous solution. ACS Nano 7, 1817–1822 (2013).

  27. 27

    Pyne, A., Thompson, R., Leung, C., Roy, D. & Hoogenboom, B. W. Single-molecule reconstruction of oligonucleotide secondary structure by atomic force microscopy. Small 10, 3257–3261 (2014).

  28. 28

    Ido, S. et al. Immunoactive two-dimensional self-assembly of monoclonal antibodies in aqueous solution revealed by atomic force microscopy. Nat. Mater. 13, 264–270 (2014).

  29. 29

    Seelert, H. et al. Proton powered turbine of a plant motor. Nature 405, 418–419 (2000).

  30. 30

    Fotiadis, D. et al. Atomic-force microscopy: rhodopsin dimers in native disc membranes. Nature 421, 127–128 (2003).

  31. 31

    Uchihashi, T., Iino, R., Ando, T. & Noji, H. High-speed atomic force microscopy reveals rotary catalysis of rotorless F1-ATPase. Science 333, 755–758 (2011).

  32. 32

    Müller, D. J., Hand, G. M., Engel, A. & Sosinsky, G. Conformational changes in surface structures of isolated Connexin26 gap junctions. EMBO J. 21, 3598–3607 (2002).

  33. 33

    Müller, D. J. & Engel, A. Voltage and pH-induced channel closure of porin OmpF visualized by atomic force microscopy. J. Mol. Biol. 285, 1347–1351 (1999).

  34. 34

    Mari, S. A. et al. pH-induced conformational change of the beta-barrel-forming protein OmpG reconstituted into native E. coli lipids. J. Mol. Biol. 396, 610–616 (2010).

  35. 35

    Czajkowsky, D. M., Hotze, E. M., Shao, Z. & Tweten, R. K. Vertical collapse of a cytolysin prepore moves its transmembrane beta-hairpins to the membrane. EMBO J. 23, 3206–3215 (2004).

  36. 36

    Shibata, M., Uchihashi, T., Yamashita, H., Kandori, H. & Ando, T. Structural changes in bacteriorhodopsin in response to alternate illumination observed by high-speed atomic force microscopy. Angew. Chem. Int. Ed. 50, 4410–4413 (2011).

  37. 37

    Mari, S. A. et al. Gating of the MlotiK1 potassium channel involves large rearrangements of the cyclic nucleotide-binding domains. Proc. Natl Acad. Sci. USA 108, 20802–20807 (2011).

  38. 38

    Rangl, M. et al. Real-time visualization of conformational changes within single MloK1 cyclic nucleotide-modulated channels. Nat. Commun. 7, 12789 (2016).

  39. 39

    Müller, D. J. et al. Observing membrane protein diffusion at subnanometer resolution. J. Mol. Biol. 327, 925–930 (2003).

  40. 40

    Karner, A. et al. Tuning membrane protein mobility by confinement into nanodomains. Nat. Nanotechnol. http://dx.doi.org/10.1038/nnano.2016.236 (2016).

  41. 41

    Kodera, N., Yamamoto, D., Ishikawa, R. & Ando, T. Video imaging of walking myosin V by high-speed atomic force microscopy. Nature 468, 72–76 (2010). This study describes single-motor proteins walking along actin filaments.

  42. 42

    Cisneros, D. A., Hung, C., Franz, C. M. & Müller, D. J. Observing growth steps of collagen self-assembly by time-lapse high-resolution atomic force microscopy. J. Struct. Biol. 154, 232–245 (2006).

  43. 43

    Stamov, D. R., Stock, E., Franz, C. M., Jahnke, T. & Haschke, H. Imaging collagen type I fibrillogenesis with high spatiotemporal resolution. Ultramicroscopy 149, 86–94 (2015).

  44. 44

    Lehto, T., Miaczynska, M., Zerial, M., Müller, D. J. & Severin, F. Observing the growth of individual actin filaments in cell extracts by time-lapse atomic force microscopy. FEBS Lett. 551, 25–28 (2003).

  45. 45

    Sharma, S. et al. Nanostructured self-assembly of inverted formin 2 (INF2) and F-actin-INF2 complexes revealed by atomic force microscopy. Langmuir 30, 7533–7539 (2014).

  46. 46

    Friedrichs, J., Taubenberger, A., Franz, C. M. & Müller, D. J. Cellular remodelling of individual collagen fibrils visualized by time-lapse AFM. J. Mol. Biol. 372, 594–607 (2007).

  47. 47

    Gudzenko, T. & Franz, C. M. Studying early stages of fibronectin fibrillogenesis in living cells by atomic force microscopy. Mol. Biol. Cell 26, 3190–3204 (2015).

  48. 48

    Stark, M., Stark, R. W., Heckl, W. M. & Guckenberger, R. Inverting dynamic force microscopy: from signals to time-resolved interaction forces. Proc. Natl Acad. Sci. USA 99, 8473–8478 (2002).

  49. 49

    Martinez-Martin, D., Herruzo, E. T., Dietz, C., Gomez-Herrero, J. & Garcia, R. Noninvasive protein structural flexibility mapping by bimodal dynamic force microscopy. Phys. Rev. Lett. 106, 198101 (2011).

  50. 50

    Raman, A. et al. Mapping nanomechanical properties of live cells using multi-harmonic atomic force microscopy. Nat. Nanotechnol. 6, 809–814 (2011).

  51. 51

    Hansma, P. K., Schitter, G., Fantner, G. E. & Prater, C. High-speed atomic force microscopy. Science 314, 601–602 (2006).

  52. 52

    Fantner, G. E. et al. Components for high speed atomic force microscopy. Ultramicroscopy 106, 881–887 (2006).

  53. 53

    Viani, M. B. et al. Probing protein–protein interactions in real time. Nat. Struct. Biol. 7, 644–647 (2000). This paper introduces the use of ultrashort AFM cantilevers to detect fast interactions, which is the basis for high-speed AFM imaging and force spectroscopy.

  54. 54

    Watanabe-Nakayama, T., Itami, M., Kodera, N., Ando, T. & Konno, H. High-speed atomic force microscopy reveals strongly polarized movement of clostridial collagenase along collagen fibrils. Sci. Rep. 6, 28975 (2016).

  55. 55

    Chiaruttini, N. et al. Relaxation of loaded ESCRT-III spiral springs drives membrane deformation. Cell 163, 866–879 (2015).

  56. 56

    Yamashita, H. et al. Single-molecule imaging on living bacterial cell surface by high-speed AFM. J. Mol. Biol. 422, 300–309 (2012).

  57. 57

    Sakiyama, Y., Mazur, A., Kapinos, L. E. & Lim, R. Y. Spatiotemporal dynamics of the nuclear pore complex transport barrier resolved by high-speed atomic force microscopy. Nat. Nanotechnol. 11, 719–723 (2016).

  58. 58

    Butt, H. J., Cappella, B. & Kappl, M. Force measurements with the atomic force microscope: technique, interpretation and applications. Surf. Sci. Rep. 59, 1–152 (2005).

  59. 59

    Ducker, W. A., Senden, T. J. & Pashley, R. M. Direct measurement of colloidal forces using an atomic force microscope. Nature 353, 239–241 (1991).

  60. 60

    Pelling, A. E., Sehati, S., Gralla, E. B., Valentine, J. S. & Gimzewski, J. K. Local nanomechanical motion of the cell wall of Saccharomyces cerevisiae. Science 305, 1147–1150 (2004).

  61. 61

    Krieg, M., Dunn, A. R. & Goodman, M. B. Mechanical control of the sense of touch by beta-spectrin. Nat. Cell Biol. 16, 224–233 (2014).

  62. 62

    Vasquez, V., Krieg, M., Lockhead, D. & Goodman, M. B. Phospholipids that contain polyunsaturated fatty acids enhance neuronal cell mechanics and touch sensation. Cell Rep. 6, 70–80 (2014).

  63. 63

    Krieg, M. et al. Tensile forces govern germ-layer organization in zebrafish. Nat. Cell Biol. 10, 429–436 (2008). This study uses AFM to settle a long-standing dispute regarding whether it is cell adhesion or cortex tension that is responsible for cell sorting in the developing zebrafish embyro.

  64. 64

    Strilic, B. et al. Electrostatic cell-surface repulsion initiates lumen formation in developing blood vessels. Curr. Biol. 20, 2003–2009 (2010).

  65. 65

    Matzke, R., Jacobson, K. & Radmacher, M. Direct, high-resolution measurement of furrow stiffening during division of adherent cells. Nat. Cell Biol. 3, 607–610 (2001).

  66. 66

    Stewart, M. P. et al. Hydrostatic pressure and the actomyosin cortex drive mitotic cell rounding. Nature 469, 226–230 (2011).

  67. 67

    Cross, S. E., Jin, Y. S., Rao, J. & Gimzewski, J. K. Nanomechanical analysis of cells from cancer patients. Nat. Nanotechnol. 2, 780–783 (2007).

  68. 68

    Iyer, S., Gaikwad, R. M., Subba-Rao, V., Woodworth, C. D. & Sokolov, I. Atomic force microscopy detects differences in the surface brush of normal and cancerous cells. Nat. Nanotechnol. 4, 389–393 (2009).

  69. 69

    Martinez-Martin, D. et al. Resolving structure and mechanical properties at the nanoscale of viruses with frequency modulation atomic force microscopy. PLoS ONE 7, e30204 (2012).

  70. 70

    Roos, W. H. et al. Mechanics of bacteriophage maturation. Proc. Natl Acad. Sci. USA 109, 2342–2347 (2012).

  71. 71

    Marchetti, M., Wuite, G. & Roos, W. H. Atomic force microscopy observation and characterization of single virions and virus-like particles by nano-indentation. Curr. Opin. Virol. 18, 82–88 (2016).

  72. 72

    Janmey, P. A., Georges, P. C. & Hvidt, S. Basic rheology for biologists. Methods Cell Biol. 83, 3–27 (2007).

  73. 73

    Nawaz, S. et al. Cell visco-elasticity measured with AFM and optical trapping at sub-micrometer deformations. PLoS ONE 7, e45297 (2012).

  74. 74

    Medalsy, I. D. & Müller, D. J. Nanomechanical properties of proteins and membranes depend on loading rate and electrostatic interactions. ACS Nano 7, 2642–2650 (2013).

  75. 75

    Herruzo, E. T., Perrino, A. P. & Garcia, R. Fast nanomechanical spectroscopy of soft matter. Nat. Commun. 5, 3126 (2014).

  76. 76

    Stewart, M. P. et al. Wedged AFM-cantilevers for parallel plate cell mechanics. Methods 60, 186–194 (2013).

  77. 77

    Fischer-Friedrich, E., Hyman, A. A., Julicher, F., Müller, D. J. & Helenius, J. Quantification of surface tension and internal pressure generated by single mitotic cells. Sci. Rep. 4, 6213 (2014).

  78. 78

    Fischer-Friedrich, E. et al. Rheology of the active cell cortex in mitosis. Biophys. J. 111, 589–600 (2016).

  79. 79

    Lee, G. U., Kidwell, D. A. & Colton, R. J. Sensing discrete streptavidin–biotin interactions with atomic force microscopy. Langmuir 10, 354–357 (1994).

  80. 80

    Moy, V. T., Florin, E. L. & Gaub, H. E. Intermolecular forces and energies between ligands and receptors. Science 266, 257–259 (1994).

  81. 81

    Baumann, F., Heucke, S. F., Pippig, D. A. & Gaub, H. E. Tip localization of an atomic force microscope in transmission microscopy with nanoscale precision. Rev. Sci. Instrum. 86, 035109 (2015).

  82. 82

    Evans, E. A. & Calderwood, D. A. Forces and bond dynamics in cell adhesion. Science 316, 1148–1153 (2007).

  83. 83

    Dudko, O. K., Hummer, G. & Szabo, A. Theory, analysis, and interpretation of single-molecule force spectroscopy experiments. Proc. Natl Acad. Sci. USA 105, 15755–15760 (2008).

  84. 84

    Friddle, R. W., Noy, A. & De Yoreo, J. J. Interpreting the widespread nonlinear force spectra of intermolecular bonds. Proc. Natl Acad. Sci. USA 109, 13573–13578 (2012).

  85. 85

    Woodside, M. T. & Block, S. M. Reconstructing folding energy landscapes by single-molecule force spectroscopy. Ann. Rev. Biophys. 43, 19–39 (2014).

  86. 86

    Perez-Jimenez, R. et al. Single-molecule paleoenzymology probes the chemistry of resurrected enzymes. Nat. Struct. Mol. Biol. 18, 592–596 (2011).

  87. 87

    Oberhauser, A. F., Hansma, P. K., Carrion-Vazquez, M. & Fernandez, J. M. Stepwise unfolding of titin under force-clamp atomic force microscopy. Proc. Natl Acad. Sci. USA 98, 468–472 (2001).

  88. 88

    Stahl, S. W., Puchner, E. M. & Gaub, H. E. Photothermal cantilever actuation for fast single-molecule force spectroscopy. Rev. Sci. Instrum. 80, 073702 (2009).

  89. 89

    Krieg, M., Helenius, J., Heisenberg, C. P. & Müller, D. J. A bond for a lifetime: employing membrane nanotubes from living cells to determine receptor–ligand kinetics. Angew. Chem. Int. Ed. 47, 9775–9777 (2008).

  90. 90

    Alsteens, D. et al. Imaging G protein-coupled receptors while quantifying their ligand-binding free-energy landscape. Nat. Methods 12, 845–851 (2015).

  91. 91

    Wildling, L. et al. Probing binding pocket of serotonin transporter by single molecular force spectroscopy on living cells. J. Biol. Chem. 287, 105–113 (2012).

  92. 92

    Friedrichs, J., Helenius, J. & Müller, D. J. Quantifying cellular adhesion to extracellular matrix components by single-cell force spectroscopy. Nat. Protoc. 5, 1353–1361 (2010).

  93. 93

    Schoeler, C. et al. Mapping mechanical force propagation through biomolecular complexes. Nano Lett. 15, 7370–7376 (2015).

  94. 94

    Sieben, C. et al. Influenza virus binds its host cell using multiple dynamic interactions. Proc. Natl Acad. Sci. USA 109, 13626–13631 (2012).

  95. 95

    Alsteens, D. et al. Nanomechanical mapping of first binding steps of a virus to animal cells. Nat. Nanotechnol. 12, 177–183 (2017). This paper maps virus-binding events on animal cells and simultaneously extracts the steps and free energy landscape of viral ligands binding to cell surface receptors.

  96. 96

    King, G. M., Carter, A. R., Churnside, A. B., Eberle, L. S. & Perkins, T. T. Ultrastable atomic force microscopy: atomic-scale stability and registration in ambient conditions. Nano Lett. 9, 1451–1456 (2009).

  97. 97

    Bull, M. S., Sullan, R. M., Li, H. & Perkins, T. T. Improved single molecule force spectroscopy using micromachined cantilevers. ACS Nano 8, 4984–4995 (2014).

  98. 98

    Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J. M. & Gaub, H. E. Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 276, 1109–1112 (1997). This paper characterizes the mechanically induced unfolding and reversible refolding of single protein domains using AFM-based SMFS.

  99. 99

    Bippes, C. A. & Müller, D. J. High-resolution atomic force microscopy and spectroscopy of native membrane proteins. Rep. Prog. Phys. 74, 086601 (2011).

  100. 100

    Puchner, E. M. & Gaub, H. E. Force and function: probing proteins with AFM-based force spectroscopy. Curr. Opin. Struct. Biol. 19, 605–614 (2009).

  101. 101

    Žoldák, G. & Rief, M. Force as a single molecule probe of multidimensional protein energy landscapes. Curr. Opin. Struct. Biol. 23, 48–57 (2013).

  102. 102

    Kawamura, S., Colozo, A. T., Ge, L., Müller, D. J. & Park, P. S. Structural, energetic, and mechanical perturbations in rhodopsin mutant that causes congenital stationary night blindness. J. Biol. Chem. 287, 21826–21835 (2012).

  103. 103

    Mashaghi, A. et al. Reshaping of the conformational search of a protein by the chaperone trigger factor. Nature 500, 98–101 (2013).

  104. 104

    Nunes, J. M., Mayer-Hartl, M., Hartl, F. U. & Müller, D. J. Action of the Hsp70 chaperone system observed with single proteins. Nat. Commun. 6, 6307 (2015).

  105. 105

    Park, P. S. et al. Stabilizing effect of Zn2+ in native bovine rhodopsin. J. Biol. Chem. 282, 11377–11385 (2007).

  106. 106

    Oesterhelt, F. et al. Unfolding pathways of individual bacteriorhodopsins. Science 288, 143–146 (2000). This paper describes the mechanically induced stepwise unfolding of membrane proteins using AFM-based SMFS.

  107. 107

    Damaghi, M., Koster, S., Bippes, C. A., Yildiz, O. & Müller, D. J. One β-hairpin follows the other: exploring refolding pathways and kinetics of the transmembrane β-barrel protein OmpG. Angew. Chem. Int. Ed. 50, 7422–7424 (2011).

  108. 108

    Kessler, M., Gottschalk, K. E., Janovjak, H., Müller, D. J. & Gaub, H. E. Bacteriorhodopsin folds into the membrane against an external force. J. Mol. Biol. 357, 644–654 (2006).

  109. 109

    Thoma, J., Bosshart, P., Pfreundschuh, M. & Müller, D. J. Out but not in: the large transmembrane β-barrel protein FhuA unfolds but cannot refold via β-hairpins. Structure 20, 2185–2190 (2012).

  110. 110

    Serdiuk, T. et al. YidC assists the stepwise and stochastic folding of membrane proteins. Nat. Chem. Biol. 12, 911–917 (2016).

  111. 111

    Thoma, J., Burmann, B. M., Hiller, S. & Müller, D. J. Impact of holdase chaperones Skp and SurA on the folding of beta-barrel outer-membrane proteins. Nat. Struct. Mol. Biol. 22, 795–802 (2015).

  112. 112

    Struckmeier, J. et al. Fully automated single-molecule force spectroscopy for screening applications. Nanotechnology 19, 384020 (2008).

  113. 113

    Otten, M. et al. From genes to protein mechanics on a chip. Nat. Methods 11, 1127–1130 (2014).

  114. 114

    Friedrichs, J. et al. A practical guide to quantify cell adhesion using single-cell force spectroscopy. Methods 60, 169–178 (2013).

  115. 115

    Benoit, M., Gabriel, D., Gerisch, G. & Gaub, H. E. Discrete interactions in cell adhesion measured by single-molecule force spectroscopy. Nat. Cell Biol. 2, 313–317 (2000). This paper introduces AFM-based force spectroscopy to quantify the adhesive forces established by living cells.

  116. 116

    Ulrich, F. et al. Wnt11 functions in gastrulation by controlling cell cohesion through Rab5c and E-cadherin. Dev. Cell 9, 555–564 (2005).

  117. 117

    Te Riet, J. et al. Dynamic coupling of ALCAM to the actin cortex strengthens cell adhesion to CD6. J. Cell Sci. 127, 1595–1606 (2014).

  118. 118

    Alsteens, D., Van Dijck, P., Lipke, P. N. & Dufrêne, Y. F. Quantifying the forces driving cell–cell adhesion in a fungal pathogen. Langmuir 29, 13473–13480 (2013).

  119. 119

    Beaussart, A. et al. Single-cell force spectroscopy of probiotic bacteria. Biophys. J. 104, 1886–1892 (2013).

  120. 120

    Friedrichs, J., Helenius, J. & Müller, D. J. Stimulated single-cell force spectroscopy to quantify cell adhesion receptor crosstalk. Proteomics 10, 1455–1462 (2010).

  121. 121

    Chaudhuri, O., Parekh, S. H., Lam, W. A. & Fletcher, D. A. Combined atomic force microscopy and side-view optical imaging for mechanical studies of cells. Nat. Methods 6, 383–387 (2009).

  122. 122

    Gonnermann, C. et al. Quantitating membrane bleb stiffness using AFM force spectroscopy and an optical sideview setup. Integr. Biol. (Camb.) 7, 356–363 (2015).

  123. 123

    Ramanathan, S. P. et al. Cdk1-dependent mitotic enrichment of cortical myosin II promotes cell rounding against confinement. Nat. Cell Biol. 17, 148–159 (2015).

  124. 124

    Rouven Bruckner, B., Pietuch, A., Nehls, S., Rother, J. & Janshoff, A. Ezrin is a major regulator of membrane tension in epithelial cells. Sci. Rep. 5, 14700 (2015).

  125. 125

    Heu, C., Berquand, A., Elie-Caille, C. & Nicod, L. Glyphosate-induced stiffening of HaCaT keratinocytes, a Peak Force Tapping study on living cells. J. Struct. Biol. 178, 1–7 (2012).

  126. 126

    Hecht, F. M. et al. Imaging viscoelastic properties of live cells by AFM: power-law rheology on the nanoscale. Soft Matter 11, 4584–4591 (2015).

  127. 127

    Formosa-Dague, C., Speziale, P., Foster, T. J., Geoghegan, J. A. & Dufrêne, Y. F. Zinc-dependent mechanical properties of Staphylococcus aureus biofilm-forming surface protein SasG. Proc. Natl Acad. Sci. USA 113, 410–415 (2016).

  128. 128

    Beaussart, A., El- Kirat-Chatel, S., Fontaine, T., Latge, J. P. & Dufrêne, Y. F. Nanoscale biophysical properties of the cell surface galactosaminogalactan from the fungal pathogen Aspergillus fumigatus. Nanoscale 7, 14996–15004 (2015).

  129. 129

    Dong, M., Husale, S. & Sahin, O. Determination of protein structural flexibility by microsecond force spectroscopy. Nat. Nanotechnol. 4, 514–517 (2009).

  130. 130

    Medalsy, I., Hensen, U. & Müller, D. J. Imaging and quantifying chemical and physical properties of native proteins at molecular resolution by force–volume AFM. Angew. Chem. Int. Ed. 50, 12103–12108 (2011).

  131. 131

    Wegmann, S., Medalsy, I. D., Mandelkow, E. & Müller, D. J. The fuzzy coat of pathological human Tau fibrils is a two-layered polyelectrolyte brush. Proc. Natl Acad. Sci. USA 110, E313–E321 (2013).

  132. 132

    Zhang, S. et al. Coexistence of ribbon and helical fibrils originating from hIAPP(20–29) revealed by quantitative nanomechanical atomic force microscopy. Proc. Natl Acad. Sci. USA 110, 2798–2803 (2013).

  133. 133

    Grandbois, M., Dettmann, W., Benoit, M. & Gaub, H. E. Affinity imaging of red blood cells using an atomic force microscope. J. Histochem. Cytochem. 48, 719–724 (2000).

  134. 134

    Dague, E. et al. Chemical force microscopy of single live cells. Nano Lett. 7, 3026–3030 (2007).

  135. 135

    Dupres, V. et al. The yeast Wsc1 cell surface sensor behaves like a nanospring in vivo. Nat. Chem. Biol. 5, 857–862 (2009).

  136. 136

    Guo, S. F. et al. Measuring protein isoelectric points by AFM-based force spectroscopy using trace amounts of sample. Nat. Nanotechnol. 11, 817–823 (2016).

  137. 137

    Pfreundschuh, M., Hensen, U. & Müller, D. J. Quantitative imaging of the electrostatic field and potential generated by a transmembrane protein pore at subnanometer resolution. Nano Lett. 13, 5585–5593 (2013).

  138. 138

    Alsteens, D., Trabelsi, H., Soumillion, P. & Dufrêne, Y. F. Multiparametric atomic force microscopy imaging of single bacteriophages extruding from living bacteria. Nat. Commun. 4, 2926 (2013).

  139. 139

    Pfreundschuh, M. et al. Identifying and quantifying two ligand-binding sites while imaging native human membrane receptors by AFM. Nat. Commun. 6, 8857 (2015).

  140. 140

    Pfreundschuh, M., Alsteens, D., Hilbert, M., Steinmetz, M. O. & Müller, D. J. Localizing chemical groups while imaging single native proteins by high-resolution atomic force microscopy. Nano Lett. 14, 2957–2964 (2014).

  141. 141

    Kim, D. & Sahin, O. Imaging and three-dimensional reconstruction of chemical groups inside a protein complex using atomic force microscopy. Nat. Nanotechnol. 10, 264–269 (2015).

  142. 142

    Dong, M. & Sahin, O. A nanomechanical interface to rapid single-molecule interactions. Nat. Commun. 2, 247 (2011).

  143. 143

    Thomas, W. E., Vogel, V. & Sokurenko, E. Biophysics of catch bonds. Ann. Rev. Biophys. 37, 399–416 (2008).

  144. 144

    Janovjak, H., Struckmeier, J. & Müller, D. J. Hydrodynamic effects in fast AFM single-molecule force measurements. Eur. Biophys. J. 34, 91–96 (2005).

  145. 145

    Amo, C. A. & Garcia, R. Fundamental high-speed limits in single-molecule, single-cell, and nanoscale force spectroscopies. ACS Nano 10, 7117–7124 (2016).

  146. 146

    Rico, F., Gonzalez, L., Casuso, I., Puig-Vidal, M. & Scheuring, S. High-speed force spectroscopy unfolds titin at the velocity of molecular dynamics simulations. Science 342, 741–743 (2013).

  147. 147

    Fritz, J. et al. Translating biomolecular recognition into nanomechanics. Science 288, 316–318 (2000). This study introduces the use of microcantilevers to sense biomolecular binding.

  148. 148

    McKendry, R. et al. Multiple label-free biodetection and quantitative DNA-binding assays on a nanomechanical cantilever array. Proc. Natl Acad. Sci. USA 99, 9783–9788 (2002).

  149. 149

    Zhang, J. et al. Rapid and label-free nanomechanical detection of biomarker transcripts in human RNA. Nat. Nanotechnol. 1, 214–220 (2006).

  150. 150

    Braun, T. et al. Quantitative time-resolved measurement of membrane protein–ligand interactions using microcantilever array sensors. Nat. Nanotechnol. 4, 179–185 (2009).

  151. 151

    Ndieyira, J. W. et al. Surface-stress sensors for rapid and ultrasensitive detection of active free drugs in human serum. Nat. Nanotechnol. 9, 225–232 (2014).

  152. 152

    Patil, S. B. et al. Decoupling competing surface binding kinetics and reconfiguration of receptor footprint for ultrasensitive stress assays. Nat. Nanotechnol. 10, 899–907 (2015).

  153. 153

    Huber, F., Lang, H. P., Backmann, N., Rimoldi, D. & Gerber, C. Direct detection of a BRAF mutation in total RNA from melanoma cells using cantilever arrays. Nat. Nanotechnol. 8, 125–129 (2013).

  154. 154

    Huber, F. et al. Fast diagnostics of BRAF mutations in biopsies from malignant melanoma. Nano Lett. 16, 5373–5377 (2016).

  155. 155

    Barnes, J. R., Stephenson, R. J., Welland, M. E., Gerber, C. & Gimzewski, J. K. Photothermal spectroscopy with femtojoule sensitivity using a micromechanical device. Nature 372, 79–81 (1994).

  156. 156

    Kasas, S. et al. Detecting nanoscale vibrations as signature of life. Proc. Natl Acad. Sci. USA 112, 378–381 (2015).

  157. 157

    Carbonell, C. & Braunschweig, A. B. Toward 4D nanoprinting with tip-induced organic surface reactions. Acc. Chem. Res. http://dx.doi.org/10.1021/acs.accounts.6b00307 (2016).

  158. 158

    Tinazli, A., Piehler, J., Beuttler, M., Guckenberger, R. & Tampe, R. Native protein nanolithography that can write, read and erase. Nat. Nanotechnol. 2, 220–225 (2007).

  159. 159

    Martinez, R. V. et al. Large-scale nanopatterning of single proteins used as carriers of magnetic nanoparticles. Adv. Mater. 22, 588–591 (2010).

  160. 160

    Felts, J. R., Onses, M. S., Rogers, J. A. & King, W. P. Nanometer scale alignment of block-copolymer domains by means of a scanning probe tip. Adv. Mater. 26, 2999–3002 (2014).

  161. 161

    Shi, J., Chen, J. & Cremer, P. S. Sub-100 nm patterning of supported bilayers by nanoshaving lithography. J. Am. Chem. Soc. 130, 2718–2719 (2008).

  162. 162

    Cisneros, D. A., Friedrichs, J., Taubenberger, A., Franz, C. M. & Müller, D. J. Creating ultrathin nanoscopic collagen matrices for biological and biotechnological applications. Small 3, 956–963 (2007).

  163. 163

    Szoszkiewicz, R. et al. High-speed, sub-15 nm feature size thermochemical nanolithography. Nano Lett. 7, 1064–1069 (2007).

  164. 164

    Paul, P. C., Knoll, A. W., Holzner, F., Despont, M. & Duerig, U. Rapid turnaround scanning probe nanolithography. Nanotechnology 22, 275306 (2011).

  165. 165

    Gotsmann, B., Duerig, U., Frommer, J. & Hawker, C. J. Exploiting chemical switching in a Diels–Alder polymer for nanoscale probe lithography and data storage. Adv. Funct. Mater. 16, 1499–1505 (2006).

  166. 166

    Milner, A. A., Zhang, K. & Prior, Y. Floating tip nanolithography. Nano Lett. 8, 2017–2022 (2008).

  167. 167

    Carroll, K. M. et al. Fabricating nanoscale chemical gradients with ThermoChemical NanoLithography. Langmuir 29, 8675–8682 (2013).

  168. 168

    Jaschke, M. et al. The atomic force microscope as a tool to study and manipulate local surface properties. Biosens. Bioelectron. 11, 601–612 (1996).

  169. 169

    Salaita, K., Wang, Y. & Mirkin, C. A. Applications of dip-pen nanolithography. Nat. Nanotechnol. 2, 145–155 (2007).

  170. 170

    Lenhert, S., Mirkin, C. A. & Fuchs, H. In situ lipid dip-pen nanolithography under water. Scanning 32, 15–23 (2010).

  171. 171

    Kim, K. H., Moldovan, N. & Espinosa, H. D. A nanofountain probe with sub-100 nm molecular writing resolution. Small 1, 632–635 (2005).

  172. 172

    Onses, M. S. et al. Hierarchical patterns of three-dimensional block-copolymer films formed by electrohydrodynamic jet printing and self-assembly. Nat. Nanotechnol. 8, 667–675 (2013).

  173. 173

    Lenhert, S. et al. Lipid multilayer gratings. Nat. Nanotechnol. 5, 275–279 (2010).

  174. 174

    Huo, F. et al. Polymer pen lithography. Science 321, 1658–1660 (2008).

  175. 175

    Albrecht, C. et al. DNA: a programmable force sensor. Science 301, 367–370 (2003).

  176. 176

    Pippig, D. A., Baumann, F., Strackharn, M., Aschenbrenner, D. & Gaub, H. E. Protein–DNA chimeras for nano assembly. ACS Nano 8, 6551–6555 (2014).

  177. 177

    Kufer, S. K. et al. Optically monitoring the mechanical assembly of single molecules. Nat. Nanotechnol. 4, 45–49 (2009).

  178. 178

    Puchner, E. M., Kufer, S. K., Strackharn, M., Stahl, S. W. & Gaub, H. E. Nanoparticle self-assembly on a DNA-scaffold written by single-molecule cut-and-paste. Nano Lett. 8, 3692–3695 (2008).

  179. 179

    Strackharn, M., Stahl, S. W., Puchner, E. M. & Gaub, H. E. Functional assembly of aptamer binding sites by single-molecule cut-and-paste. Nano Lett. 12, 2425–2428 (2012).

  180. 180

    Franz, C. M. & Müller, D. J. Analysing focal adhesion structure by AFM. J. Cell Sci. 118, 5315–5323 (2005).

  181. 181

    Cordes, T. et al. Resolving single-molecule assembled patterns with superresolution blink-microscopy. Nano Lett. 10, 645–651 (2010).

  182. 182

    Monserrate, A., Casado, S. & Flors, C. Correlative atomic force microscopy and localization-based super-resolution microscopy: revealing labelling and image reconstruction artefacts. Chemphyschem 15, 647–650 (2014).

  183. 183

    Fukuda, S. et al. High-speed atomic force microscope combined with single-molecule fluorescence microscope. Rev. Sci. Instrum. 84, 073706 (2013).

  184. 184

    Schmid, T., Opilik, L., Blum, C. & Zenobi, R. Nanoscale chemical imaging using tip-enhanced Raman spectroscopy: a critical review. Angew. Chem. Int. Ed. 52, 5940–5954 (2013).

  185. 185

    Berweger, S. et al. Nano-chemical infrared imaging of membrane proteins in lipid bilayers. J. Am. Chem. Soc. 135, 18292–18295 (2013).

  186. 186

    Ruggeri, F. S. et al. Infrared nanospectroscopy characterization of oligomeric and fibrillar aggregates during amyloid formation. Nat. Commun. 6, 7831 (2015).

  187. 187

    Hansma, P. K., Drake, B., Marti, O., Gould, S. A. & Prater, C. B. The scanning ion-conductance microscope. Science 243, 641–643 (1989). This classic study describes the invention of scanning ion conductance microscopy.

  188. 188

    Novak, P. et al. Nanoscale live-cell imaging using hopping probe ion conductance microscopy. Nat. Methods 6, 279–281 (2009).

  189. 189

    Novak, P. et al. Nanoscale-targeted patch-clamp recordings of functional presynaptic ion channels. Neuron 79, 1067–1077 (2013).

  190. 190

    Takahashi, Y. et al. Multifunctional nanoprobes for nanoscale chemical imaging and localized chemical delivery at surfaces and interfaces. Angew. Chem. Int. Ed. 50, 9638–9642 (2011).

  191. 191

    Shevchuk, A. I. et al. An alternative mechanism of clathrin-coated pit closure revealed by ion conductance microscopy. J. Cell Biol. 197, 499–508 (2012).

  192. 192

    Novak, P. et al. Imaging single nanoparticle interactions with human lung cells using fast ion conductance microscopy. Nano Lett. 14, 1202–1207 (2014).

  193. 193

    Shevchuk, A. I. et al. Imaging proteins in membranes of living cells by high-resolution scanning ion conductance microscopy. Angew. Chem. Int. Ed. 45, 2212–2216 (2006).

  194. 194

    Klausen, L. H., Fuhs, T. & Dong, M. Mapping surface charge density of lipid bilayers by quantitative surface conductivity microscopy. Nat. Commun. 7, 12447 (2016).

  195. 195

    Ossola, D. et al. Simultaneous scanning ion conductance microscopy and atomic force microscopy with microchanneled cantilevers. Phys. Rev. Lett. 115, 238103 (2015).

  196. 196

    Leo-Macias, A. et al. Nanoscale visualization of functional adhesion/excitability nodes at the intercalated disc. Nat. Commun. 7, 10342 (2016).

  197. 197

    Galvagnion, C. et al. Lipid vesicles trigger alpha-synuclein aggregation by stimulating primary nucleation. Nat. Chem. Biol. 11, 229–234 (2015).

  198. 198

    Lind, T. K., Zielinska, P., Wacklin, H. P., Urbanczyk-Lipkowska, Z. & Cardenas, M. Continuous flow atomic force microscopy imaging reveals fluidity and time-dependent interactions of antimicrobial dendrimer with model lipid membranes. ACS Nano 8, 396–408 (2014).

  199. 199

    Ko, S. H. et al. Synergistic self-assembly of RNA and DNA molecules. Nat. Chem. 2, 1050–1055 (2010).

  200. 200

    Sapra, K. T. et al. One beta hairpin after the other: exploring mechanical unfolding pathways of the transmembrane beta-barrel protein OmpG. Angew. Chem. Int. Ed. 48, 8306–8308 (2009).

  201. 201

    Strackharn, M., Pippig, D. A., Meyer, P., Stahl, S. W. & Gaub, H. E. Nanoscale arrangement of proteins by single-molecule cut-and-paste. J. Am. Chem. Soc. 134, 15193–15196 (2012).

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Acknowledgements

D.A. was supported by the Belgian National Foundation for Scientific Research (FNRS) and the Université catholique de Louvain (Fonds Spéciaux de Recherche). D.A. is a Research Associate FNRS. D.J.M. was supported by the Swiss National Science Foundation (SNF; grant 310030B_160225) and the NCCR Molecular Systems Engineering. C.G. and D.J.M. were supported by the Swiss Nanoscience Institute. H.E.G. acknowledges financial support by the ERC grant CelluFuel.

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Alsteens, D., Gaub, H., Newton, R. et al. Atomic force microscopy-based characterization and design of biointerfaces. Nat Rev Mater 2, 17008 (2017). https://doi.org/10.1038/natrevmats.2017.8

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