Review Article | Published:

Super-resolution microscopy as a powerful tool to study complex synthetic materials

Nature Reviews Chemistryvolume 3pages6884 (2019) | Download Citation

Abstract

Understanding the relations between the formation, structure, dynamics and functionality of complex synthetic materials is one of the great challenges in chemistry and nanotechnology and represents the foundation for the rational design of novel materials for a variety of applications. Initially conceived to study biology below the diffraction limit, super-resolution microscopy (SRM) is emerging as a powerful tool for studying synthetic materials owing to its nanometric resolution, multicolour ability and minimal invasiveness. In this Review, we provide an overview of the pioneering studies that use SRM to visualize materials, highlighting exciting recent developments such as experiments in operando, wherein materials, such as biomaterials in a biological environment, are imaged in action. Moreover, the potential and the challenges of the different SRM methods for application in nanotechnology and (bio)materials science are discussed, aiming to guide researchers to select the best SRM approach for their specific purpose.

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References

  1. 1.

    Huang, B., Bates, M. & Zhuang, X. Super-resolution fluorescence microscopy. Annu. Rev. Biochem. 78, 993–1016 (2009).

  2. 2.

    Schermelleh, L., Heintzmann, R. & Leonhardt, H. A guide to super-resolution fluorescence microscopy. J. Cell Biol. 190, 165–175 (2010).

  3. 3.

    Möckl, L., Lamb, D. C. & Bräuchle, C. Super-resolved fluorescence microscopy: Nobel Prize in Chemistry 2014 for Eric Betzig, Stefan Hell, and William E. Moerner. Angew. Chem. Int. Ed. 53, 13972–13977 (2014).

  4. 4.

    Binnig, G., Quate, C. F. & Gerber, C. Atomic force microscope. Phys. Rev. Lett. 56, 930–933 (1986).

  5. 5.

    Mathys, D. Die Entwicklung der Elektronenmikroskopie vom Bild über die Analyse zum Nanolabor (University of Basel, 2004).

  6. 6.

    Webber, M. J., Appel, E. A., Meijer, E. W. & Langer, R. Supramolecular biomaterials. Nat. Mater. 15, 13–26 (2016).

  7. 7.

    Hyotyla, J. T. & Lim, R. Y. H. in Supramolecular Chemistry: from Molecules to Nanomaterials (eds Steed, J. W. & Gale, P. A.) (John Wiley & Sons, Ltd, 2012).

  8. 8.

    Franken, L. E., Boekema, E. J. & Stuart, M. C. A. Transmission electron microscopy as a tool for the characterization of soft materials: application and interpretation. Adv. Sci. 4, 1600476 (2017).

  9. 9.

    Lakowicz, J. R. Principles of Fluorescence Spectroscopy (Springer, 2011).

  10. 10.

    Douglass, K. M., Sieben, C., Archetti, A., Lambert, A. & Manley, S. Super-resolution imaging of multiple cells by optimized flat-field epi-illumination. Nat. Photon. 10, 705–708 (2016).

  11. 11.

    Bates, M., Dempsey, G. T., Chen, K. H. & Zhuang, X. Multicolor super-resolution fluorescence imaging via multi-parameter fluorophore detection. Chemphyschem 13, 99–107 (2012).

  12. 12.

    Jungmann, R. et al. Multiplexed 3D cellular super-resolution imaging with DNA-PAINT and Exchange-PAINT. Nat. Methods 11, 313–318 (2014).

  13. 13.

    Lee, S.-H., Shin, J. Y., Lee, A. & Bustamante, C. Counting single photoactivatable fluorescent molecules by photoactivated localization microscopy (PALM). Proc. Natl Acad. Sci. USA 109, 17436–17441 (2012).

  14. 14.

    Venkataramani, V., Herrmannsdörfer, F., Heilemann, M. & Kuner, T. SuReSim: simulating localization microscopy experiments from ground truth models. Nat. Methods 13, 319–321 (2016).

  15. 15.

    Habuchi, S. Super-resolution molecular and functional imaging of nanoscale architectures in life and materials science. Front. Bioeng. Biotechnol. 2, 20 (2014).

  16. 16.

    Van Loon, J., Kubarev, A. V. & Roeffaers, M. B. J. Correlating catalyst structure and activity at the nanoscale. ChemNanoMat 4, 6–14 (2018).

  17. 17.

    Aida, T., Meijer, E. W. & Stupp, S. I. Functional supramolecular polymers. Science 335, 813–817 (2012).

  18. 18.

    Albertazzi, L. et al. Probing exchange pathways in one-dimensional aggregates with super-resolution microscopy. Science 344, 491–495 (2014).

  19. 19.

    Baker, M. B. et al. Consequences of chirality on the dynamics of a water-soluble supramolecular polymer. Nat. Commun. 6, 6234 (2015).

  20. 20.

    Baker, M. B. et al. Exposing differences in monomer exchange rates of multicomponent supramolecular polymers in water. ChemBioChem 17, 207–213 (2016).

  21. 21.

    Aloi, A. et al. Imaging nanostructures by single-molecule localization microscopy in organic solvents. J. Am. Chem. Soc. 138, 2953–2956 (2016).

  22. 22.

    Boott, C. E. et al. In situ visualization of block copolymer self-assembly in organic media by super-resolution fluorescence microscopy. Chemistry 21, 18539–18542 (2015).

  23. 23.

    Adelizzi, B. et al. Painting supramolecular polymers in organic solvents by super-resolution microscopy. ACS Nano 12, 4431–4439 (2018).

  24. 24.

    Hendrikse, S. I. S. et al. Controlling and tuning the dynamic nature of supramolecular polymers in aqueous solutions. Chem. Commun. 53, 2279–2282 (2017).

  25. 25.

    Lee, O.-S., Stupp, S. I. & Schatz, G. C. Atomistic molecular dynamics simulations of peptide amphiphile self-assembly into cylindrical nanofibers. J. Am. Chem. Soc. 133, 3677–3683 (2011).

  26. 26.

    Aloi, A., Vilanova, N., Albertazzi, L. & Voets, I. K. iPAINT: a general approach tailored to image the topology of interfaces with nanometer resolution. Nanoscale 8, 8712–8716 (2016).

  27. 27.

    Merdasa, A. et al. Single Lévy states–disorder induced energy funnels in molecular aggregates. Nano Lett. 14, 6774–6781 (2014).

  28. 28.

    Onogi, S. et al. In situ real-time imaging of self-sorted supramolecular nanofibres. Nat. Chem. 8, 743–752 (2016).

  29. 29.

    Tønnesen, J., Inavalli, V. V. G. K. & Nägerl, U. V. Super-resolution imaging of the extracellular space in living brain tissue. Cell 172, 1108–1121 (2018).

  30. 30.

    Schacher, F. H., Rupar, P. A. & Manners, I. Functional block copolymers: nanostructured materials with emerging applications. Angew. Chem. Int. Ed. 51, 7898–7921 (2012).

  31. 31.

    Qiu, H. et al. Uniform patchy and hollow rectangular platelet micelles from crystallizable polymer blends. Science 352, 697–701 (2016).

  32. 32.

    Ullal, C. K., Schmidt, R., Hell, S. W. & Egner, A. Block copolymer nanostructures mapped by far-field optics. Nano Lett. 9, 2497–2500 (2009).

  33. 33.

    Yan, J. et al. Optical nanoimaging for block copolymer self-assembly. J. Am. Chem. Soc. 137, 2436–2439 (2015).

  34. 34.

    González-Aramundiz, J. V., Lozano, M. V., Sousa-Herves, A., Fernandez-Megia, E. & Csaba, N. Polypeptides and polyaminoacids in drug delivery. Expert Opin. Drug Deliv. 9, 183–201 (2012).

  35. 35.

    Beun, L. H., Albertazzi, L., van der Zwaag, D., de Vries, R. & Cohen Stuart, M. A. Unidirectional living growth of self-assembled protein nanofibrils revealed by super-resolution microscopy. ACS Nano 10, 4973–4980 (2016).

  36. 36.

    Duro-Castano, A. et al. Capturing “extraordinary” soft-assembled charge-like polypeptides as a strategy for nanocarrier design. Adv. Mater. 29, 1702888 (2017).

  37. 37.

    Muls, B. et al. Direct measurement of the end-to-end distance of individual polyfluorene polymer chains. Chemphyschem 6, 2286–2294 (2005).

  38. 38.

    Vacha, M. & Habuchi, S. Conformation and physics of polymer chains: a single-molecule perspective. NPG Asia Mater. 2, 134–142 (2010).

  39. 39.

    Aoki, H., Mori, K. & Ito, S. Conformational analysis of single polymer chains in three dimensions by super-resolution fluorescence microscopy. Soft Matter 8, 4390–4395 (2012).

  40. 40.

    Gramlich, M. W., Bae, J., Hayward, R. C. & Ross, J. L. Fluorescence imaging of nanoscale domains in polymer blends using stochastic optical reconstruction microscopy (STORM). Opt. Express 22, 8438–8450 (2014).

  41. 41.

    O.Neil, C. E., Jackson, J. M., Shim, S.-H. & Soper, S. A. Interrogating surface functional group heterogeneity of activated thermoplastics using super-resolution fluorescence microscopy. Anal. Chem. 88, 3686–3696 (2016).

  42. 42.

    Bolinger, J. C., Traub, M. C., Adachi, T. & Barbara, P. F. Ultralong-range polaron-induced quenching of excitons in isolated conjugated polymers. Science 331, 565–567 (2011).

  43. 43.

    Habuchi, S., Onda, S. & Vacha, M. Mapping the emitting sites within a single conjugated polymer molecule. Chem. Commun. 0, 4868–4870 (2009).

  44. 44.

    Habuchi, S., Onda, S. & Vacha, M. Molecular weight dependence of emission intensity and emitting sites distribution within single conjugated polymer molecules. Phys. Chem. Chem. Phys 13, 1743–1753 (2011).

  45. 45.

    Park, H., Hoang, D. T., Paeng, K. & Kaufman, L. J. Localizing exciton recombination sites in conformationally distinct single conjugated polymers by super-resolution fluorescence imaging. ACS Nano 9, 3151–3158 (2015).

  46. 46.

    Penwell, S. B., Ginsberg, L. D. S. & Ginsberg, N. S. Bringing far-field subdiffraction optical imaging to electronically coupled optoelectronic molecular materials using their endogenous chromophores. J. Phys. Chem. Lett. 6, 2767–2772 (2015).

  47. 47.

    King, J. T. & Granick, S. Operating organic light-emitting diodes imaged by super-resolution spectroscopy. Nat. Commun. 7, 11691 (2016).

  48. 48.

    Tian, Z., Li, A. D. Q. & Hu, D. Super-resolution fluorescence nanoscopy applied to imaging core–shell photoswitching nanoparticles and their self-assemblies. Chem. Commun. 47, 1258–1260 (2011).

  49. 49.

    Hu, D., Tian, Z., Wu, W., Wan, W. & Li, A. D. Q. Photoswitchable nanoparticles enable high-resolution cell imaging: PULSAR microscopy. J. Am. Chem. Soc. 130, 15279–15281 (2008).

  50. 50.

    Gong, W.-L. et al. Single-wavelength-controlled in situ dynamic super-resolution fluorescence imaging for block copolymer nanostructures via blue-light-switchable FRAP. Photochem. Photobiol. Sci. 15, 1433–1441 (2016).

  51. 51.

    Nevskyi, O., Sysoiev, D., Oppermann, A., Huhn, T. & Wöll, D. Nanoscopic visualization of soft matter using fluorescent diarylethene photoswitches. Angew. Chem. Int. Ed. 55, 12698–12702 (2016).

  52. 52.

    Urban, B. E. et al. Subsurface super-resolution imaging of unstained polymer nanostructures. Sci. Rep. 6, 28156 (2016).

  53. 53.

    Barenholz, Y. Doxil®—the first FDA-approved nano-drug: lessons learned. J. Control. Release 160, 117–134 (2012).

  54. 54.

    Sharonov, A. & Hochstrasser, R. M. Wide-field subdiffraction imaging by accumulated binding of diffusing probes. Proc. Natl Acad. Sci. USA 103, 18911–18916 (2006).

  55. 55.

    Kuo, C. & Hochstrasser, R. M. Super-resolution microscopy of lipid bilayer phases. J. Am. Chem. Soc. 133, 4664–4667 (2011).

  56. 56.

    Bongiovanni, M. N. et al. Multi-dimensional super-resolution imaging enables surface hydrophobicity mapping. Nat. Commun. 7, 13544 (2016).

  57. 57.

    Yan, R., Moon, S., Kenny, S. J. & Xu, K. Spectrally resolved and functional super-resolution microscopy via ultrahigh-throughput single-molecule spectroscopy. Acc. Chem. Res. 51, 697–705 (2018).

  58. 58.

    Boreham, A., Volz, P., Peters, D., Keck, C. M. & Alexiev, U. Determination of nanostructures and drug distribution in lipid nanoparticles by single molecule microscopy. Eur. J. Pharm. Biopharm. 110, 31–38 (2017).

  59. 59.

    Belfiore, L., Spenkelink, L. M., Ranson, M., van Oijen, A. M. & Vine, K. L. Quantification of ligand density and stoichiometry on the surface of liposomes using single-molecule fluorescence imaging. J. Control. Release 278, 80–86 (2018).

  60. 60.

    Rothemund, P. W. K. Folding DNA to create nanoscale shapes and patterns. Nature 440, 297–302 (2006).

  61. 61.

    Saccà, B. & Niemeyer, C. M. DNA origami: the art of folding DNA. Angew. Chem. Int. Ed Engl. 51, 58–66 (2012).

  62. 62.

    Graugnard, E., Hughes, W. L., Jungmann, R., Kostiainen, M. A. & Linko, V. Nanometrology and super-resolution imaging with DNA. MRS Bull. 42, 951–959 (2017).

  63. 63.

    Steinhauer, C., Jungmann, R., Sobey, T., Simmel, F. & Tinnefeld, P. DNA origami as a nanoscopic ruler for super-resolution microscopy. Angew. Chem. Int. Ed. 48, 8870–8873 (2009).

  64. 64.

    Schmied, J. J. et al. DNA origami nanopillars as standards for three-dimensional superresolution microscopy. Nano Lett. 13, 781–785 (2013).

  65. 65.

    Schmied, J. J. et al. Fluorescence and super-resolution standards based on DNA origami. Nat. Methods 9, 1133–1134 (2012).

  66. 66.

    Jungmann, R. et al. Single-molecule kinetics and super-resolution microscopy by fluorescence imaging of transient binding on DNA origami. Nano Lett. 10, 4756–4761 (2010).

  67. 67.

    Smith, D. M. et al. A structurally variable hinged tetrahedron framework from DNA origami. J. Nucleic Acids 2011, 360954 (2011).

  68. 68.

    Scheible, M. B. et al. A compact DNA cube with side length 10nm. Small 11, 5200–5205 (2015).

  69. 69.

    Chen, J., Bremauntz, A., Kisley, L., Shuang, B. & Landes, C. F. Super-resolution mbPAINT for optical localization of single-stranded DNA. ACS Appl. Mater. Interfaces 5, 9338–9343 (2013).

  70. 70.

    Knudsen, J. B. et al. Routing of individual polymers in designed patterns. Nat. Nanotechnol. 10, 892–898 (2015).

  71. 71.

    Schueder, F. et al. Multiplexed 3D super-resolution imaging of whole cells using spinning disk confocal microscopy and DNA-PAINT. Nat. Commun. 8, 2090 (2017).

  72. 72.

    Auer, A., Strauss, M. T., Schlichthaerle, T. & Jungmann, R. Fast, background-free DNA-PAINT imaging using FRET-based probes. Nano Lett. 17, 6428–6434 (2017).

  73. 73.

    Karathanasis, C., Fricke, F., Hummer, G. & Heilemann, M. Molecule counts in localization microscopy with organic fluorophores. Chemphyschem 18, 942–948 (2017).

  74. 74.

    Beater, S., Holzmeister, P., Pibiri, E., Lalkens, B. & Tinnefeld, P. Choosing dyes for cw-STED nanoscopy using self-assembled nanorulers. Phys. Chem. Chem. Phys 16, 6990–6996 (2014).

  75. 75.

    Dai, M., Jungmann, R. & Yin, P. Optical imaging of individual biomolecules in densely packed clusters. Nat. Nanotechnol. 11, 798–807 (2016).

  76. 76.

    Delcanale, P., Miret-Ontiveros, B., Arista-Romero, M., Pujals, S. & Albertazzi, L. Nanoscale mapping functional sites on nanoparticles by points accumulation for imaging in nanoscale topography (PAINT). ACS Nano 12, 7629–7637 (2018).

  77. 77.

    Harke, B., Ullal, C. K., Keller, J. & Hell, S. W. Three-dimensional nanoscopy of colloidal crystals. Nano Lett. 8, 1309–1313 (2008).

  78. 78.

    Hauser, M. et al. Correlative super-resolution microscopy: new dimensions and new opportunities. Chem. Rev. 117, 7428–7456 (2017).

  79. 79.

    Bon, P. et al. Three-dimensional nanometre localization of nanoparticles to enhance super-resolution microscopy. Nat. Commun. 6, 7764 (2015).

  80. 80.

    Xu, W., Kong, J. S., Yeh, Y.-T. E. & Chen, P. Single-molecule nanocatalysis reveals heterogeneous reaction pathways and catalytic dynamics. Nat. Mater. 7, 992–996 (2008).

  81. 81.

    Chen, T., Zhang, Y. & Xu, W. Single-molecule nanocatalysis reveals catalytic activation energy of single nanocatalysts. J. Am. Chem. Soc. 138, 12414–12421 (2016).

  82. 82.

    Zhou, X., Choudhary, E., Andoy, N. M., Zou, N. & Chen, P. Scalable parallel screening of catalyst activity at the single-particle level and subdiffraction resolution. ACS Catal. 3, 1448–1453 (2013).

  83. 83.

    Zhou, X. et al. Quantitative super-resolution imaging uncovers reactivity patterns on single nanocatalysts. Nat. Nanotechnol. 7, 237–241 (2012).

  84. 84.

    Cang, H. et al. Probing the electromagnetic field of a 15-nanometre hotspot by single molecule imaging. Nature 469, 385–388 (2011).

  85. 85.

    Xu, W., Kong, J. S. & Chen, P. Probing the catalytic activity and heterogeneity of Au-nanoparticles at the single-molecule level. Phys. Chem. Chem. Phys 11, 2767–2778 (2009).

  86. 86.

    Zhou, X., Xu, W., Liu, G., Panda, D. & Chen, P. Size-dependent catalytic activity and dynamics of gold nanoparticles at the single-molecule level. J. Am. Chem. Soc. 132, 138–146 (2010).

  87. 87.

    Chen, T., Zhang, Y. & Xu, W. Size-dependent catalytic kinetics and dynamics of Pd nanocubes: a single-particle study. Phys. Chem. Chem. Phys 18, 22494–22502 (2016).

  88. 88.

    Shen, H., Zhou, X., Zou, N. & Chen, P. Single-molecule kinetics reveals a hidden surface reaction intermediate in single-nanoparticle catalysis. J. Phys. Chem. C 118, 26902–26911 (2014).

  89. 89.

    Han, K. S., Liu, G., Zhou, X., Medina, R. E. & Chen, P. How does a single Pt nanocatalyst behave in two different reactions? a single-molecule study. Nano Lett. 12, 1253–1259 (2012).

  90. 90.

    Wilson, A. J., Molina, N. Y. & Willets, K. A. Modification of the electrochemical properties of nile blue through covalent attachment to gold as revealed by electrochemistry and SERS. J. Phys. Chem. C 120, 21091–21098 (2016).

  91. 91.

    Wilson, A. J. & Willets, K. A. Unforeseen distance-dependent SERS spectroelectrochemistry from surface-tethered Nile Blue: the role of molecular orientation. Analyst 141, 5144–5151 (2016).

  92. 92.

    Titus, E. J., Weber, M. L., Stranahan, S. M. & Willets, K. A. Super-resolution SERS imaging beyond the single-molecule limit: an isotope-edited approach. Nano Lett. 12, 5103–5110 (2012).

  93. 93.

    Willets, K. A., Wilson, A. J., Sundaresan, V. & Joshi, P. B. Super-resolution imaging and plasmonics. Chem. Rev. 117, 7538–7582 (2017).

  94. 94.

    Blythe, K. L., Mayer, K. M., Weber, M. L. & Willets, K. A. Ground state depletion microscopy for imaging interactions between gold nanowires and fluorophore-labeled ligands. Phys. Chem. Chem. Phys 15, 4136–4145 (2013).

  95. 95.

    Lim, K. et al. Nanostructure-induced distortion in single-emitter microscopy. Nano Lett. 16, 5415–5419 (2016).

  96. 96.

    Thompson, R. E., Larson, D. R. & Webb, W. W. Precise nanometer localization analysis for individual fluorescent probes. Biophys. J. 82, 2775–2783 (2002).

  97. 97.

    Lin, H. et al. Mapping of surface-enhanced fluorescence on metal nanoparticles using super-resolution photoactivation localization microscopy. Chemphyschem 13, 973–981 (2012).

  98. 98.

    Simoncelli, S., Roberti, M. J., Araoz, B., Bossi, M. L. & Aramendía, P. F. Mapping the fluorescence performance of a photochromic–fluorescent system coupled with gold nanoparticles at the single-molecule–single-particle level. J. Am. Chem. Soc. 136, 6878–6880 (2014).

  99. 99.

    Fu, B., Flynn, J. D., Isaacoff, B. P., Rowland, D. J. & Biteen, J. S. Super-resolving the distance-dependent plasmon-enhanced fluorescence of single dye and fluorescent protein molecules. J. Phys. Chem. C 119, 19350–19358 (2015).

  100. 100.

    Johlin, E. et al. Super-resolution imaging of light–matter interactions near single semiconductor nanowires. Nat. Commun. 7, 13950 (2016).

  101. 101.

    Simoncelli, S., Li, Y., Cortés, E. & Maier, S. A. Nanoscale control of molecular self-assembly induced by plasmonic hot-electron dynamics. ACS Nano 12, 2184–2192 (2018).

  102. 102.

    Taylor, A., Verhoef, R., Beuwer, M., Wang, Y. & Zijlstra, P. All-optical imaging of gold nanoparticle geometry using super-resolution microscopy. J. Phys. Chem. C Nanomater. Interfaces 122, 2336–2342 (2018).

  103. 103.

    Yuan, H. et al. Imaging heterogeneously distributed photo-active traps in perovskite single crystals. Adv. Mater. 30, 1705494 (2018).

  104. 104.

    Ristanovic´, Z. et al. High-resolution single-molecule fluorescence imaging of zeolite aggregates within real-life fluid catalytic cracking particles. Angew. Chem. Int. Ed. 54, 1836–1840 (2015).

  105. 105.

    Ristanovic´, Z. et al. Quantitative 3D fluorescence imaging of single catalytic turnovers reveals spatiotemporal gradients in reactivity of zeolite H-ZSM-5 crystals upon steaming. J. Am. Chem. Soc. 137, 6559–6568 (2015).

  106. 106.

    Roeffaers, M. B. J. et al. Super-resolution reactivity mapping of nanostructured catalyst particles. Angew. Chem. Int. Ed. 48, 9285–9289 (2009).

  107. 107.

    Hendriks, F. C. et al. Integrated transmission electron and single-molecule fluorescence microscopy correlates reactivity with ultrastructure in a single catalyst particle. Angew. Chem. Int. Ed. 57, 257–261 (2018).

  108. 108.

    Roeffaers, M. B. J. et al. Spatially resolved observation of crystal-face-dependent catalysis by single turnover counting. Nature 439, 572–575 (2006).

  109. 109.

    Cavalieri, F. et al. Redox-sensitive PEG–polypeptide nanoporous particles for survivin silencing in prostate cancer cells. Biomacromolecules 16, 2168–2178 (2015).

  110. 110.

    Teplensky, M. H. et al. Temperature treatment of highly porous zirconium-containing metal–organic frameworks extends drug delivery release. J. Am. Chem. Soc. 139, 7522–7532 (2017).

  111. 111.

    Tolstik, E. et al. Studies of silicon nanoparticles uptake and biodegradation in cancer cells by Raman spectroscopy. Nanomedicine 12, 1931–1940 (2016).

  112. 112.

    Tolstik, E. et al. Linear and non-linear optical imaging of cancer cells with silicon nanoparticles. Int. J. Mol. Sci. 17, E1536 (2016).

  113. 113.

    Guggenheim, E. J. et al. Comparison of confocal and super-resolution reflectance imaging of metal oxide nanoparticles. PLOS ONE 11, e0159980 (2016).

  114. 114.

    Schübbe, S. et al. Size-dependent localization and quantitative evaluation of the intracellular migration of silica nanoparticles in caco-2 cells. Chem. Mater. 24, 914–923 (2012).

  115. 115.

    Schübbe, S., Cavelius, C., Schumann, C., Koch, M. & Kraegeloh, A. STED microscopy to monitor agglomeration of silica particles inside A549 cells. Adv. Eng. Mater. 12, 417–422 (2010).

  116. 116.

    Peuschel, H., Ruckelshausen, T., Cavelius, C. & Kraegeloh, A. Quantification of internalized silica nanoparticles via STED microscopy. Biomed Res. Int. 2015, 961208 (2015).

  117. 117.

    Leménager, G., Luca, E. D., Sun, Y.-P. & Pompa, P. P. Super-resolution fluorescence imaging of biocompatible carbon dots. Nanoscale 6, 8617–8623 (2014).

  118. 118.

    Shang, L. et al. Protein-based fluorescent nanoparticles for super-resolution STED imaging of live cells. Chem. Sci. 8, 2396–2400 (2017).

  119. 119.

    Tzeng, Y.-K. et al. Superresolution imaging of albumin-conjugated fluorescent nanodiamonds in cells by stimulated emission depletion. Angew. Chem. Int. Ed. 50, 2262–2265 (2011).

  120. 120.

    Wäldchen, S., Lehmann, J., Klein, T., van de Linde, S. & Sauer, M. Light-induced cell damage in live-cell super-resolution microscopy. Sci. Rep. 5, 15348 (2015).

  121. 121.

    Heine, J. et al. Adaptive-illumination STED nanoscopy. Proc. Natl Acad. Sci. USA 114, 9797–9802 (2017).

  122. 122.

    Göttfert, F. et al. Strong signal increase in STED fluorescence microscopy by imaging regions of subdiffraction extent. Proc. Natl Acad. Sci. USA 114, 2125–2130 (2017).

  123. 123.

    van der Zwaag, D. et al. Super resolution imaging of nanoparticles cellular uptake and trafficking. ACS Appl. Mater. Interfaces 8, 6391–6399 (2016).

  124. 124.

    De Koker, S. et al. Engineering polymer hydrogel nanoparticles for lymph node-targeted delivery. Angew. Chem. Int. Ed. 55, 1334–1339 (2016).

  125. 125.

    Li, Y., Shang, L. & Nienhaus, G. U. Super-resolution imaging-based single particle tracking reveals dynamics of nanoparticle internalization by live cells. Nanoscale 8, 7423–7429 (2016).

  126. 126.

    Geddes, C. D., Parfenov, A., Gryczynski, I. & Lakowicz, J. R. Luminescent blinking of gold nanoparticles. Chem. Phys. Lett. 380, 269–272 (2003).

  127. 127.

    Kuno, M., Fromm, D. P., Hamann, H. F., Gallagher, A. & Nesbitt, D. J. “On”/“off” fluorescence intermittency of single semiconductor quantum dots. J. Chem. Phys. 115, 1028–1040 (2001).

  128. 128.

    Moser, F. et al. Cellular uptake of gold nanoparticles and their behavior as labels for localization microscopy. Biophys. J. 110, 947–953 (2016).

  129. 129.

    Wegner, W. et al. In vivo mouse and live cell STED microscopy of neuronal actin plasticity using far-red emitting fluorescent proteins. Sci. Rep. 7, 11781 (2017).

  130. 130.

    Wegner, W., Mott, A. C., Grant, S. G. N., Steffens, H. & Willig, K. I. In vivo STED microscopy visualizes PSD95 sub-structures and morphological changes over several hours in the mouse visual cortex. Sci. Rep. 8, 219 (2018).

  131. 131.

    Cui, J. et al. Immobilized particle imaging for quantification of nano- and microparticles. Langmuir 32, 3532–3540 (2016).

  132. 132.

    Ardizzone, A. et al. Nanostructuring lipophilic dyes in water using stable vesicles, quatsomes, as scaffolds and their use as probes for bioimaging. Small 14, 1703851 (2018).

  133. 133.

    Krivitsky, A. et al. Amphiphilic poly(α)glutamate polymeric micelles for systemic administration of siRNA to tumors. Nanomedicine 14, 303–315 (2018).

  134. 134.

    Chen, X. et al. Analysing intracellular deformation of polymer capsules using structured illumination microscopy. Nanoscale 8, 11924–11931 (2016).

  135. 135.

    Chen, X. et al. Probing cell internalisation mechanics with polymer capsules. Nanoscale 8, 17096–17101 (2016).

  136. 136.

    Wilhelm, S. et al. Analysis of nanoparticle delivery to tumours. Nat. Rev. Mater. 1, 16014 (2016).

  137. 137.

    Ke, P. C., Lin, S., Parak, W. J., Davis, T. P. & Caruso, F. A. Decade of the protein corona. ACS Nano 11, 11773–11776 (2017).

  138. 138.

    Feiner-Gracia, N. et al. Super-resolution microscopy unveils dynamic heterogeneities in nanoparticle protein corona. Small 13, 1701631 (2017).

  139. 139.

    Clemments, A. M., Botella, P. & Landry, C. C. Spatial mapping of protein adsorption on mesoporous silica nanoparticles by stochastic optical reconstruction microscopy. J. Am. Chem. Soc. 139, 3978–3981 (2017).

  140. 140.

    Runa, S., Lakadamyali, M., Kemp, M. L. & Payne, C. K. TiO2 nanoparticle-induced oxidation of the plasma membrane: importance of the protein corona. J. Phys. Chem. B 121, 8619–8625 (2017).

  141. 141.

    Oria, R. et al. Force loading explains spatial sensing of ligands by cells. Nature 552, 219–224 (2017).

  142. 142.

    Chelladurai, R., Debnath, K., Jana, N. R. & Basu, J. K. Nanoscale heterogeneities drive enhanced binding and anomalous diffusion of nanoparticles in model biomembranes. Langmuir 34, 1691–1699 (2018).

  143. 143.

    Chmyrov, A. et al. Nanoscopy with more than 100,000 ‘doughnuts’. Nat. Methods 10, 737–740 (2013).

  144. 144.

    Winter, P. W. et al. Incoherent structured illumination improves optical sectioning and contrast in multiphoton super-resolution microscopy. Opt. Express 23, 5327–5334 (2015).

  145. 145.

    Izeddin, I. et al. PSF shaping using adaptive optics for three-dimensional single-molecule super-resolution imaging and tracking. Opt. Express 20, 4957–4967 (2012).

  146. 146.

    Uno, S. et al. A spontaneously blinking fluorophore based on intramolecular spirocyclization for live-cell super-resolution imaging. Nat. Chem. 6, 681–689 (2014).

  147. 147.

    Hsiao, W. W.-W., Hui, Y. Y., Tsai, P.-C. & Chang, H.-C. Fluorescent nanodiamond: a versatile tool for long-term cell tracking, super-resolution imaging, and nanoscale temperature sensing. Acc. Chem. Res. 49, 400–407 (2016).

  148. 148.

    Kianinia, M. et al. All-optical control and super-resolution imaging of quantum emitters in layered materials. Nat. Commun. 9, 874 (2018).

  149. 149.

    Betzig, E. et al. Imaging intracellular fluorescent proteins at nanometer resolution. Science 313, 1642–1645 (2006).

  150. 150.

    Watanabe, S. et al. Protein localization in electron micrographs using fluorescence nanoscopy. Nat. Methods 8, 80–84 (2011).

  151. 151.

    Kopek, B. G., Shtengel, G., Xu, C. S., Clayton, D. A. & Hess, H. F. Correlative 3D superresolution fluorescence and electron microscopy reveal the relationship of mitochondrial nucleoids to membranes. Proc. Natl Acad. Sci. USA 109, 6136–6141 (2012).

  152. 152.

    Kopek, B. G., Shtengel, G., Grimm, J. B., Clayton, D. A. & Hess, H. F. Correlative photoactivated localization and scanning electron microscopy. PLOS ONE 8, e77209 (2013).

  153. 153.

    Suleiman, H. et al. Nanoscale protein architecture of the kidney glomerular basement membrane. eLife 2, e01149 (2013).

  154. 154.

    Sochacki, K. A., Shtengel, G., van Engelenburg, S. B., Hess, H. F. & Taraska, J. W. Correlative super-resolution fluorescence and metal-replica transmission electron microscopy. Nat. Methods 11, 305–308 (2014).

  155. 155.

    Engelenburg, S. B. V. et al. Distribution of ESCRT machinery at HIV assembly sites reveals virus scaffolding of ESCRT subunits. Science 343, 653–656 (2014).

  156. 156.

    Perkovic, M. et al. Correlative light- and electron microscopy with chemical tags. J. Struct. Biol. 186, 205–213 (2014).

  157. 157.

    Chang, Y.-W. et al. Correlated cryogenic photoactivated localization microscopy and cryo-electron tomography. Nat. Methods 11, 737–739 (2014).

  158. 158.

    Löschberger, A., Franke, C., Krohne, G., van de Linde, S. & Sauer, M. Correlative super-resolution fluorescence and electron microscopy of the nuclear pore complex with molecular resolution. J. Cell Sci. 127, 4351–4355 (2014).

  159. 159.

    Jord, A. A. et al. Centriole amplification by mother and daughter centrioles differs in multiciliated cells. Nature 516, 104–107 (2014).

  160. 160.

    Paez-Segala, M. G. et al. Fixation-resistant photoactivatable fluorescent proteins for CLEM. Nat. Methods 12, 215–218 (2015).

  161. 161.

    Johnson, E. et al. Correlative in-resin super-resolution and electron microscopy using standard fluorescent proteins. Sci. Rep. 5, 9583 (2015).

  162. 162.

    Kim, D. et al. Correlative stochastic optical reconstruction microscopy and electron microscopy. PLOS ONE 10, e0124581 (2015).

  163. 163.

    Wojcik, M., Hauser, M., Li, W., Moon, S. & Xu, K. Graphene-enabled electron microscopy and correlated super-resolution microscopy of wet cells. Nat. Commun. 6, 7384 (2015).

  164. 164.

    Liu, B. et al. Three-dimensional super-resolution protein localization correlated with vitrified cellular context. Sci. Rep. 5, 13017 (2015).

  165. 165.

    Harke, B., Chacko, J. V., Haschke, H., Canale, C. & Diaspro, A. A novel nanoscopic tool by combining AFM with STED microscopy. Opt. Nanosc. 1, 3 (2012).

  166. 166.

    Odermatt, P. D. et al. High-resolution correlative microscopy: bridging the gap between single molecule localization microscopy and atomic force microscopy. Nano Lett. 15, 4896–4904 (2015).

  167. 167.

    Chacko, J. V., Canale, C., Harke, B. & Diaspro, A. Sub-diffraction nano manipulation using STED AFM. PLOS ONE 8, e66608 (2013).

  168. 168.

    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 (2013).

  169. 169.

    Chacko, J. V., Zanacchi, F. C. & Diaspro, A. Probing cytoskeletal structures by coupling optical superresolution and AFM techniques for a correlative approach. Cytoskeleton (Hoboken) 70, 729–740 (2013).

  170. 170.

    Frasconi, M. et al. Multi-functionalized carbon nano-onions as imaging probes for cancer cells. Chemistry 21, 19071–19080 (2015).

  171. 171.

    Fenaroli, F. et al. Nanoparticles as drug delivery system against tuberculosis in zebrafish embryos: direct visualization and treatment. ACS Nano 8, 7014–7026 (2014).

  172. 172.

    Othman, B. A. et al. Correlative light-electron microscopy shows RGD-targeted ZnO nanoparticles dissolve in the intracellular environment of triple negative breast cancer cells and cause apoptosis with intratumor heterogeneity. Adv. Healthc. Mater. 5, 1310–1325 (2016).

  173. 173.

    Reifarth, M. et al. Cellular uptake of PLA nanoparticles studied by light and electron microscopy: synthesis, characterization and biocompatibility studies using an iridium(III) complex as correlative label. Chem. Commun. 52, 4361–4364 (2016).

  174. 174.

    Kempen, P. J. et al. A correlative optical microscopy and scanning electron microscopy approach to locating nanoparticles in brain tumors. Micron 68, 70–76 (2015).

  175. 175.

    Gustafsson, M. G. L. Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy. J. Microsc. 198, 82–87 (2000).

  176. 176.

    Godin, A. G., Lounis, B. & Cognet, L. Super-resolution microscopy approaches for live cell imaging. Biophys. J. 107, 1777–1784 (2014).

  177. 177.

    Gustafsson, M. G. L. Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution. Proc. Natl Acad. Sci. USA 102, 13081–13086 (2005).

  178. 178.

    York, A. G. et al. Instant super-resolution imaging in live cells and embryos via analog image processing. Nat. Methods 10, 1122–1126 (2013).

  179. 179.

    Hell, S. W. & Wichmann, J. Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy. Opt. Lett. 19, 780–782 (1994).

  180. 180.

    Klar, T. A., Jakobs, S., Dyba, M., Egner, A. & Hell, S. W. Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission. Proc. Natl Acad. Sci. USA 97, 8206–8210 (2000).

  181. 181.

    Gustafsson, M. G. L. et al. Three-dimensional resolution doubling in wide-field fluorescence microscopy by structured illumination. Biophys. J. 94, 4957–4970 (2008).

  182. 182.

    Dyba, M. & Hell, S. W. Focal spots of size λ / 23 open up far-field florescence microscopy at 33 nm axial resolution. Phys. Rev. Lett. 88, 163901 (2002).

  183. 183.

    Hess, S. T., Girirajan, T. P. K. & Mason, M. D. Ultra-high resolution imaging by fluorescence photoactivation localization microscopy. Biophys. J. 91, 4258–4272 (2006).

  184. 184.

    Rust, M. J., Bates, M. & Zhuang, X. Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat. Methods 3, 793–796 (2006).

  185. 185.

    Jones, S. A., Shim, S.-H., He, J. & Zhuang, X. Fast, three-dimensional super-resolution imaging of live cells. Nat. Methods 8, 499–505 (2011).

  186. 186.

    York, A. G., Ghitani, A., Vaziri, A., Davidson, M. W. & Shroff, H. Confined activation and subdiffractive localization enables whole-cell PALM with genetically expressed probes. Nat. Methods 8, 327–333 (2011).

  187. 187.

    Heilemann, M. et al. Subdiffraction-resolution fluorescence imaging with conventional fluorescent probes. Angew. Chem. Int. Ed. 47, 6172–6176 (2008).

  188. 188.

    Huang, B., Wang, W., Bates, M. & Zhuang, X. Three-dimensional super-resolution imaging by stochastic optical reconstruction microscopy. Science 319, 810–813 (2008).

  189. 189.

    Lin, C. et al. Submicrometre geometrically encoded fluorescent barcodes self-assembled from DNA. Nat. Chem. 4, 832–839 (2012).

  190. 190.

    Iinuma, R. et al. Polyhedra self-assembled from DNA tripods and characterized with 3D DNA-PAINT. Science 344, 65–69 (2014).

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Acknowledgements

The authors thank the Spanish Ministry of Economy, Industry and Competitiveness through the project SAF2016-75241-R, the Generalitat de Catalunya through the Centres de Recerca de Catalunya (CERCA) programme, the EuroNanoMed II platform through the NanoVax project, the Obra Social La Caixa foundation and the European Research Council (ERC-StG-757397). The useful discussions with the entire Nanoscopy for Nanomedicine group are gratefully acknowledged.

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Affiliations

  1. Institute for Bioengineering of Catalonia (IBEC), The Barcelona Institute of Science and Technology (BIST), Barcelona, Spain

    • Silvia Pujals
    • , Natalia Feiner-Gracia
    • , Pietro Delcanale
    •  & Lorenzo Albertazzi
  2. Laboratory of Self-Organizing Soft Matter, Laboratory of Macromolecular and Organic Chemistry, Department of Chemical Engineering, Institute for Complex Molecular Systems (ICMS), Eindhoven University of Technology, Eindhoven, Netherlands

    • Ilja Voets
  3. Department of Biomedical Engineering, Institute for Complex Molecular Systems (ICMS), Eindhoven University of Technology, Eindhoven, Netherlands

    • Lorenzo Albertazzi

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The authors contributed equally to the article.

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The authors declare no competing interests.

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Correspondence to Lorenzo Albertazzi.

Glossary

Abbe’s criteria

Criteria that state the resolution that can be obtained in principle, considering the diffraction of light. Ernst Abbe described in 1873 that by using light with a wavelength λ travelling through a medium of refractive index n and focused with a half-angle θ, the minimum resolution possible is λ/2nsin θ.

Far-field optical techniques

Techniques that make use of optical microscopes in which light does not pass through subwavelength features.

4Pi microscopy

A fluorescence technique in which two objective lenses are focused to the same spatial location to achieve improved axial resolution.

Recombinant proteins

Translated products of the expression of recombinant DNA non-native to living cells (such as bacteria, mammalian and yeast).

Stepwise photobleaching

The sequential loss of the fluorescence of individual molecules, resulting in a series of distinguishable steps that provide information about the number of fluorophores present.

Transmission diffraction grating

A device with a periodic structure that diffracts incident light into different directions.

Protein corona

The layer of adsorbed protein on the surface of a nanoparticle exposed to biological fluids such as blood. It can be divided into hard corona, stably bound proteins and soft corona, which is loosely and reversibly bound to the surface.

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https://doi.org/10.1038/s41570-018-0070-2