Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Article
  • Published:

Monitoring protein conformational changes using fluorescent nanoantennas

Nature Methods volume 19, pages 71–80 (2022)Cite this article

Subjects

Abstract

Understanding the relationship between protein structural dynamics and function is crucial for both basic research and biotechnology. However, methods for studying the fast dynamics of structural changes are limited. Here, we introduce fluorescent nanoantennas as a spectroscopic technique to sense and report protein conformational changes through noncovalent dye-protein interactions. Using experiments and molecular simulations, we detect and characterize five distinct conformational states of intestinal alkaline phosphatase, including the transient enzyme–substrate complex. We also explored the universality of the nanoantenna strategy with another model protein, Protein G and its interaction with antibodies, and demonstrated a rapid screening strategy to identify efficient nanoantennas. These versatile nanoantennas can be used with diverse dyes to monitor small and large conformational changes, suggesting that they could be used to characterize diverse protein movements or in high-throughput screening applications.

Access through your institution
Buy or subscribe

This is a preview of subscription content, access via your institution

Access options

Access through your institution

Buy this article

  • Purchase on SpringerLink
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: Overview of the fluorescent nanoantenna strategy to probe different regions on a protein.
Fig. 2: Fluorescent nanoantennas enable complete characterization of the enzyme’s kinetic mechanism.
Fig. 3: Fluorescent nanoantennas enabled real-time monitoring of any substrate hydrolyzed by AP.
Fig. 4: Screening inhibitors by monitoring AP hydrolysis using fluorescent nanoantennas.
Fig. 5: Detecting other states of AP using fluorescent nanoantennas.
Fig. 6: Rapid screening strategy for identification of functional nanoantennas for protein G binding IgG.

Similar content being viewed by others

Data availability

The fluorescence kinetic signatures of the substrates and inhibitors have been deposited on figshare (https://doi.org/10.6084/m9.figshare.16798174).

Code availability

The MATLAB fitting script for substrate kinetics is provided in the Supplementary Information.

References

  1. Hammes-Schiffer, S. & Klinman, J. Emerging concepts about the role of protein motion in enzyme catalysis. Acc. Chem. Res. 48, 899–899 (2015).

    Article  CAS  PubMed  Google Scholar 

  2. Lobb, R. R. & Auld, D. S. Determination of enzyme mechanisms by radiationless energy transfer kinetics. Proc. Natl Acad. Sci. USA 76, 2684–2688 (1979).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Au, H.-W., Tsang, M.-W., So, P.-K., Wong, K.-Y. & Leung, Y.-C. Thermostable β-lactamase mutant with its active site conjugated with fluorescein for efficient β-lactam antibiotic detection. ACS Omega 4, 20493–20502 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Vallée-Bélisle, A. & Michnick, S. W. Visualizing transient protein-folding intermediates by tryptophan-scanning mutagenesis. Nat. Struct. Mol. Biol. 19, 731–736 (2012).

    Article  PubMed  Google Scholar 

  5. Gregorio, G. G. et al. Single-molecule analysis of ligand efficacy in β2AR–G-protein activation. Nature 547, 68–73 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  6. Comstock, M. J. et al. Direct observation of structure-function relationship in a nucleic acid–processing enzyme. Science 348, 352–354 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Hwang, H. & Myong, S. Protein induced fluorescence enhancement (PIFE) for probing protein–nucleic acid interactions. Chem. Soc. Rev. 43, 1221–1229 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. Lerner, E. et al. Toward dynamic structural biology: two decades of single-molecule Förster resonance energy transfer. Science 359, eaan1133 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  9. Chen, Y., Tsao, K. & Keillor, J. W. Fluorogenic protein labelling: a review of photophysical quench mechanisms and principles of fluorogen design. Can. J. Chem. 93, 389–398 (2015).

    Article  CAS  Google Scholar 

  10. Unnikrishnan, B., Wu, R.-S., Wei, S.-C., Huang, C.-C. & Chang, H.-T. Fluorescent carbon dots for selective labeling of subcellular organelles. ACS Omega 5, 11248–11261 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  11. Eisenmesser, E. Z., Bosco, D. A., Akke, M. & Kern, D. Enzyme dynamics during catalysis. Science 295, 1520–1523 (2002).

    Article  CAS  PubMed  Google Scholar 

  12. Ma, X., Hortelão, A. C., Patiño, T. & Sánchez, S. Enzyme catalysis to power micro/nanomachines. ACS Nano 10, 9111–9122 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Abou-Zied, O. K. & Sulaiman, S. A. J. Site-specific recognition of fluorescein by human serum albumin: a steady-state and time-resolved spectroscopic study. Dyes Pigment. 110, 89–96 (2014).

    Article  CAS  Google Scholar 

  14. Pisoni, D. S. et al. Symmetrical and asymmetrical cyanine dyes. Synthesis, spectral properties, and BSA association study. J. Org. Chem. 79, 5511–5520 (2014).

    Article  CAS  PubMed  Google Scholar 

  15. Millán, J.L. Mammalian Alkaline Phosphatases: From Biology to Applications in Medicine and Biotechnology (John Wiley & Sons, 2006).

  16. Lallès, J.-P. Recent advances in intestinal alkaline phosphatase, inflammation, and nutrition. Nutr. Rev. 77, 710–724 (2019).

    Article  PubMed  Google Scholar 

  17. Bates, J. M., Akerlund, J., Mittge, E. & Guillemin, K. Intestinal alkaline phosphatase detoxifies lipopolysaccharide and prevents inflammation in zebrafish in response to the gut microbiota. Cell Host Microbe 2, 371–382 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Malo, M. S. et al. Intestinal alkaline phosphatase promotes gut bacterial growth by reducing the concentration of luminal nucleotide triphosphates. Am. J. Physiol. Gastrointest. Liver Physiol. 306, G826–G838 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Mizumori, M. et al. Intestinal alkaline phosphatase regulates protective surface microclimate pH in rat duodenum. J. Physiol. 587, 3651–3663 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Giatromanolaki, A., Sivridis, E., Maltezos, E. & Koukourakis, M. I. Down-regulation of intestinal-type alkaline phosphatase in the tumor vasculature and stroma provides a strong basis for explaining amifostine selectivity. Semin. Oncol. 29, 14–21 (2002).

    Article  CAS  PubMed  Google Scholar 

  21. Hofer, M. et al. Two new faces of amifostine: protector from DNA damage in normal cells and inhibitor of DNA repair in cancer cells. J. Med. Chem. 59, 3003–3017 (2016).

    Article  CAS  PubMed  Google Scholar 

  22. Riedel, C. et al. The heat released during catalytic turnover enhances the diffusion of an enzyme. Nature 517, 227–230 (2015).

    Article  CAS  PubMed  Google Scholar 

  23. Tsai, L.-C. et al. Expression and regulation of alkaline phosphatases in human breast cancer MCF-7 cells. Eur. J. Biochem. 267, 1330–1339 (2000).

    Article  CAS  PubMed  Google Scholar 

  24. Rao, S. R. et al. Tumour-derived alkaline phosphatase regulates tumour growth, epithelial plasticity and disease-free survival in metastatic prostate cancer. Br. J. Cancer 116, 227–236 (2017).

    Article  CAS  PubMed  Google Scholar 

  25. Hung, H.-Y. et al. Preoperative alkaline phosphatase elevation was associated with poor survival in colorectal cancer patients. Int. J. Colorectal Dis. 32, 1775–1778 (2017).

    Article  PubMed  Google Scholar 

  26. Namikawa, T. et al. Prognostic significance of serum alkaline phosphatase and lactate dehydrogenase levels in patients with unresectable advanced gastric cancer. Gastric Cancer 22, 684–691 (2019).

    Article  CAS  PubMed  Google Scholar 

  27. Kaliannan, K. et al. Intestinal alkaline phosphatase prevents metabolic syndrome in mice. Proc. Natl Acad. Sci. USA 110, 7003–7008 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. José, L. M. & Michael, P. W. Alkaline Phosphatase and Hypophosphatasia. Calcif. Tissue Int. 98, 398–416 (2016).

    Article  Google Scholar 

  29. Waymire, K. G. et al. Mice lacking tissue non-specific alkaline phosphatase die from seizures due to defective metabolism of vitamin B-6. Nat. Genet. 11, 45–51 (1995).

    Article  CAS  PubMed  Google Scholar 

  30. Park, J.-B. et al. Serum alkaline phosphatase is a predictor of mortality, myocardial infarction, or stent thrombosis after implantation of coronary drug-eluting stent. Eur. Heart J. 34, 920–931 (2012).

    Article  PubMed  Google Scholar 

  31. Yang, W. H. et al. Recurrent infection progressively disables host protection against intestinal inflammation. Science 358, eaao5610 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  32. To, K. K.-W. et al. Temporal profiles of viral load in posterior oropharyngeal saliva samples and serum antibody responses during infection by SARS-CoV-2: an observational cohort study. Lancet Infect. Dis. 20, 565–574 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Stec, B., Holtz, K. M. & Kantrowitz, E. R. A revised mechanism for the alkaline phosphatase reaction involving three metal ions. J. Mol. Biol. 299, 1303–1311 (2000).

    Article  CAS  PubMed  Google Scholar 

  34. Holtz, K. M., Stec, B. & Kantrowitz, E. R. A model of the transition state in the alkaline phosphatase reaction. J. Biol. Chem. 274, 8351–8354 (1999).

    Article  CAS  PubMed  Google Scholar 

  35. Peck, A., Sunden, F., Andrews, L. D., Pande, V. S. & Herschlag, D. Tungstate as a transition state analog for catalysis by alkaline phosphatase. J. Mol. Biol. 428, 2758–2768 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  36. Roston, D., Demapan, D. & Cui, Q. Leaving group ability observably affects transition state structure in a single enzyme active site. J. Am. Chem. Soc. 138, 7386–7394 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Bortolato, M., Besson, F. & Roux, B. Role of metal ions on the secondary and quaternary structure of alkaline phosphatase from bovine intestinal mucosa. Proteins 37, 310–318 (1999).

    Article  CAS  PubMed  Google Scholar 

  38. Ásgeirsson, B., Markússon, S., Hlynsdóttir, S. S., Helland, R. & Hjörleifsson, J. G. X-ray crystal structure of Vibrio alkaline phosphatase with the non-competitive inhibitor cyclohexylamine. Biochem. Biophys. Rep. 24, 100830 (2020).

    PubMed  PubMed Central  Google Scholar 

  39. Aziz, H. et al. Synthesis, characterization, in vitro tissue-nonspecific alkaline phosphatase (TNAP) and intestinal alkaline phosphatase (IAP) inhibition studies and computational evaluation of novel thiazole derivatives. Bioorg. Chem. 102, 104088 (2020).

    Article  CAS  PubMed  Google Scholar 

  40. Kiffer-Moreira, T. et al. Catalytic signature of a heat-stable, chimeric human alkaline phosphatase with therapeutic potential. PLoS ONE 9, e89374 (2014).

    Article  PubMed  PubMed Central  Google Scholar 

  41. Jiang, Y., Li, X. & Walt, D. R. Single-molecule analysis determines isozymes of human alkaline phosphatase in serum. Angew. Chem. Int. Ed. Engl. 59, 18010–18015 (2020).

    Article  CAS  PubMed  Google Scholar 

  42. Bessey, O. A., Lowry, O. H. & Brock, M. J. A method for the rapid determination of alkaline phosphatase with five cubic millimeters of serum. J. Biol. Chem. 164, 321–329 (1946).

    Article  CAS  PubMed  Google Scholar 

  43. Fernley, H. & Walker, P. Kinetic behaviour of calf-intestinal alkaline phosphatase with 4-methylumbelliferyl phosphate. Biochem. J. 97, 95–103 (1965).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Deng, J., Yu, P., Wang, Y. & Mao, L. Real-time ratiometric fluorescent assay for alkaline phosphatase activity with stimulus responsive infinite coordination polymer nanoparticles. Anal. Chem. 87, 3080–3086 (2015).

    Article  CAS  PubMed  Google Scholar 

  45. Sanzhaeva, U. et al. Imaging of enzyme activity by electron paramagnetic resonance: concept and experiment using a paramagnetic substrate of alkaline phosphatase. Angew. Chem. Int. Ed. Engl. 57, 11701–11705 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Gyurcsányi, R. E., Bereczki, A., Nagy, G., Neuman, M. R. & Lindner, E. Amperometric microcells for alkaline phosphatase assay. Analyst 127, 235–240 (2002).

    Article  PubMed  Google Scholar 

  47. Baykov, A. A., Evtushenko, O. A. & Avaeva, S. M. A malachite green procedure for orthophosphate determination and its use in alkaline phosphatase-based enzyme immunoassay. Anal. Biochem. 171, 266–270 (1988).

    Article  CAS  PubMed  Google Scholar 

  48. Liu, Y. & Schanze, K. S. Conjugated polyelectrolyte-based real-time fluorescence assay for alkaline phosphatase with pyrophosphate as substrate. Anal. Chem. 80, 8605–8612 (2008).

    Article  CAS  PubMed  Google Scholar 

  49. Liu, Y. et al. Selective sensing of phosphorylated peptides and monitoring kinase and phosphatase activity with a supramolecular tandem assay. J. Am. Chem. Soc. 140, 13869–13877 (2018).

    Article  CAS  PubMed  Google Scholar 

  50. Wang, Y., Wang, G., Moitessier, N. & Mittermaier, A. K. Enzyme kinetics by isothermal titration calorimetry: allostery, inhibition, and dynamics. Front. Mol. Biosci. 7, 583826 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Di Trani, J. M., Moitessier, N. & Mittermaier, A. K. Complete kinetic characterization of enzyme inhibition in a single isothermal titration calorimetric experiment. Anal. Chem. 90, 8430–8435 (2018).

    Article  PubMed  Google Scholar 

  52. Honarmand Ebrahimi, K., Hagedoorn, P.-L., Jacobs, D. & Hagen, W. R. Accurate label-free reaction kinetics determination using initial rate heat measurements. Sci. Rep. 5, 16380 (2015).

    Article  CAS  Google Scholar 

  53. Zhang, L., Buchet, R. & Azzar, G. Phosphate binding in the active site of alkaline phosphatase and the interactions of 2-nitrosoacetophenone with alkaline phosphatase-induced small structural changes. Biophys. J. 86, 3873–3881 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  54. Akerström, B., Brodin, T., Reis, K. & Björck, L. Protein G: a powerful tool for binding and detection of monoclonal and polyclonal antibodies. J. Immunol. 135, 2589–2592 (1985).

    Article  PubMed  Google Scholar 

  55. Kada, G., Falk, H. & Gruber, H. J. Accurate measurement of avidin and streptavidin in crude biofluids with a new, optimized biotin–fluorescein conjugate. Biochim. Biophys. Acta 1427, 33–43 (1999).

    Article  CAS  PubMed  Google Scholar 

  56. Buranda, T. et al. Ligand receptor dynamics at streptavidin-coated particle surfaces: a flow cytometric and spectrofluorimetric study. J. Phys. Chem. B 103, 3399–3410 (1999).

    Article  CAS  Google Scholar 

  57. Iyer, A., Chandra, A. & Swaminathan, R. Hydrolytic enzymes conjugated to quantum dots mostly retain whole catalytic activity. Biochim. Biophys. Acta Gen. Subj. 1840, 2935–2943 (2014).

    Article  CAS  Google Scholar 

  58. Fairhead, M., Krndija, D., Lowe, E. D. & Howarth, M. Plug-and-play pairing via defined divalent streptavidins. J. Mol. Biol. 426, 199–214 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. Neish, C. S., Martin, I. L., Henderson, R. M. & Edwardson, J. M. Direct visualization of ligand-protein interactions using atomic force microscopy. Br. J. Pharmacol. 135, 1943–1950 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Deetanya, P. et al. Interaction of 8-anilinonaphthalene-1-sulfonate with SARS-CoV-2 main protease and its application as a fluorescent probe for inhibitor identification. Comput. Struct. Biotechnol. J. 19, 3364–3371 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Chen, H., Ahsan, S. S., Santiago-Berrios, M. E. B., Abruña, H. D. & Webb, W. W. Mechanisms of quenching of Alexa fluorophores by natural amino acids. J. Am. Chem. Soc. 132, 7244–7245 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  62. Togashi, D. M., Szczupak, B., Ryder, A. G., Calvet, A. & O’Loughlin, M. Investigating tryptophan quenching of fluorescein fluorescence under protolytic equilibrium. J. Phys. Chem. A 113, 2757–2767 (2009).

    Article  CAS  PubMed  Google Scholar 

  63. Nguyen, B., Ciuba, M. A., Kozlov, A. G., Levitus, M. & Lohman, T. M. Protein environment and DNA orientation affect protein-induced Cy3 fluorescence enhancement. Biophys. J. 117, 66–73 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Rashid, F. et al. Initial state of DNA-Dye complex sets the stage for protein induced fluorescence modulation. Nat. Commun. 10, 2104 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  65. Marras, S. A. E., Kramer, F. R. & Tyagi, S. Efficiencies of fluorescence resonance energy transfer and contact‐mediated quenching in oligonucleotide probes. Nucleic Acids Res. 30, e122 (2002).

    Article  PubMed  PubMed Central  Google Scholar 

  66. Zimmerle, C. T. & Frieden, C. Analysis of progress curves by simulations generated by numerical integration. Biochem. J. 258, 381–387 (1989).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  67. Palmier, M. O. & Van Doren, S. R. Rapid determination of enzyme kinetics from fluorescence: overcoming the inner filter effect. Anal. Biochem. 371, 43–51 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Komazin, G. et al. Substrate structure-activity relationship reveals a limited lipopolysaccharide chemotype range for intestinal alkaline phosphatase. J. Biol. Chem. 294, 19405–19423 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Ziegler, A. J., Florian, J., Ballicora, M. A. & Herlinger, A. W. Alkaline phosphatase inhibition by vanadyl-β-diketone complexes: electron density effects. J. Enzym. Inhib. Med. Chem. 24, 22–28 (2009).

    Article  CAS  Google Scholar 

  70. Chen, S. et al. Detection of dihydrofolate reductase conformational change by FRET using two fluorescent amino acids. J. Am. Chem. Soc. 135, 12924–12927 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  71. Schwaminger, S. P. et al. Immobilization of PETase enzymes on magnetic iron oxide nanoparticles for the decomposition of microplastic PET. Nanoscale Adv. 3, 4395–4399 (2021).

    Article  CAS  Google Scholar 

  72. Ritchie, R. J. & Prvan, T. A simulation study on designing experiments to measure the Km of Michaelis–Menten kinetics curves. J. Theor. Biol. 178, 239–254 (1996).

    Article  CAS  Google Scholar 

  73. Mao, H., Yang, T. & Cremer, P. S. Design and characterization of immobilized enzymes in microfluidic systems. Anal. Chem. 74, 379–385 (2002).

    Article  CAS  PubMed  Google Scholar 

  74. Gordon, S. E., Munari, M. & Zagotta, W. N. Visualizing conformational dynamics of proteins in solution and at the cell membrane. eLife 7, e37248 (2018).

    Google Scholar 

  75. Pantazis, A., Westerberg, K., Althoff, T., Abramson, J. & Olcese, R. Harnessing photoinduced electron transfer to optically determine protein sub-nanoscale atomic distances. Nat. Commun. 9, 4738 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  76. Jarecki, BrianW. et al. Tethered spectroscopic probes estimate dynamic distances with subnanometer resolution in voltage-dependent potassium channels. Biophys. J. 105, 2724–2732 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  77. Mansoor, S. E., DeWitt, M. A. & Farrens, D. L. Distance mapping in proteins using fluorescence spectroscopy: the tryptophan-induced quenching (TrIQ) method. Biochemistry 49, 9722–9731 (2010).

    Article  CAS  PubMed  Google Scholar 

  78. Perri, M. J. & Weber, S. H. Web-Based Job Submission Interface for the GAMESS Computational Chemistry Program. J. Chem. Educ. 91, 2206–2208 (2014).

    Article  CAS  Google Scholar 

  79. Grosdidier, A., Zoete, V. & Michielin, O. SwissDock, a protein-small molecule docking web service based on EADock DSS. Nucleic Acids Res. 39, W270–W277 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  80. Grosdidier, A., Zoete, V. & Michielin, O. Fast docking using the CHARMM force field with EADock DSS. J. Comput. Chem. 32, 2149–2159 (2011).

    Article  CAS  PubMed  Google Scholar 

  81. Basu, S., Finke, A., Vera, L., Wang, M. & Olieric, V. Making routine native SAD a reality: lessons from beamline X06DA at the Swiss Light Source. Acta Crystallogr. D Struct. Biol. 75, 262–271 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Weissig, H., Schildge, A., Hoylaerts, M. F., Iqbal, M. & Millán, J. L. Cloning and expression of the bovine intestinal alkaline phosphatase gene: biochemical characterization of the recombinant enzyme. Biochem. J. 290, 503–508 (1993).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  83. Llinas, P. et al. Structural studies of human placental alkaline phosphatase in complex with functional ligands. J. Mol. Biol. 350, 441–451 (2005).

    Article  CAS  PubMed  Google Scholar 

  84. Harada, T. et al. Characterization of structural and catalytic differences in rat intestinal alkaline phosphatase isozymes. FEBS J. 272, 2477–2486 (2005).

    Article  CAS  PubMed  Google Scholar 

  85. Hanwell, M. D. et al. Avogadro: an advanced semantic chemical editor, visualization, and analysis platform. J. Cheminformatics 4, 17 (2012).

    Article  CAS  Google Scholar 

  86. Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).

    Article  CAS  PubMed  Google Scholar 

  87. Molecular Operating Environment (MOE) v.2019.01 (Chemical Computing Group, 2019).

  88. Case, D. A. et al. Amber 2020 (University of California, San Francisco, 2020).

Download references

Acknowledgements

We acknowledge scholarships from the Natural Sciences and Engineering Research Council of Canada, NSERC (S.G.H., A.D.), Fonds de recherche du Québec–Nature et technologies, FRQNT (S.G.H., D.L.), Groupe de recherche universitaire sur le medicament, GRUM (S.G.H.) and Bourse du Fonds Wilrose Desrosiers et Pauline Dunn (S.G.H.). This work was funded by NSERC, grant numbers RGPIN-2020-06975, RGPIN-06403 (A.V.-B.), Canada Reserch Chairs, grant number 950-230012 (A.V.-B.) and Le regroupement québécois de recherche sur la fonction, l’ingénierie et les applications des protéines (PROTEO). We acknowledge past and present members of our laboratory for insightful discussions, particularly C. Prévost-Tremblay (Faculté de médecine, UdeM).

Author information

Authors and Affiliations

  1. Laboratoire de Biosenseurs & Nanomachines, Département de chimie, Université de Montréal, Montréal, Québec, Canada

    Scott G. Harroun, Dominic Lauzon, Arnaud Desrosiers, Xiaomeng Wang & Alexis Vallée-Bélisle

  2. Chemical Computing Group ULC, Montréal, Québec, Canada

    Maximilian C. C. J. C. Ebert

  3. Département de biochimie et médecine moléculaire, Université de Montréal, Montréal, Québec, Canada

    Arnaud Desrosiers & Alexis Vallée-Bélisle

Authors
  1. Scott G. Harroun
    View author publications

    Search author on:PubMed Google Scholar

  2. Dominic Lauzon
    View author publications

    Search author on:PubMed Google Scholar

  3. Maximilian C. C. J. C. Ebert
    View author publications

    Search author on:PubMed Google Scholar

  4. Arnaud Desrosiers
    View author publications

    Search author on:PubMed Google Scholar

  5. Xiaomeng Wang
    View author publications

    Search author on:PubMed Google Scholar

  6. Alexis Vallée-Bélisle
    View author publications

    Search author on:PubMed Google Scholar

Contributions

S.G.H. and A.V.-B. conceived and designed the study. S.G.H. performed all experiments and the molecular docking simulations. A.D. provided ideas in an early phase of the project. D.L. wrote and used the MATLAB script to extract enzyme kinetic parameters. M.C.C.J.C.E. performed molecular dynamics simulations. X.W. realized the synthesis of the nanoantenna-AP conjugate. S.G.H. and A.V.-B. created the figures and wrote the manuscript, which was then reviewed and edited by all of the authors. A.V.-B. provided project oversight and funding.

Corresponding author

Correspondence to Alexis Vallée-Bélisle.

Ethics declarations

Competing interests

The authors declare no competing interests.

Peer review information

Nature Methods thanks the anonymous reviewers for their contribution to the peer review of this work. Rita Strack was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data

Extended Data Fig. 1 Progress of a typical enzymatic reaction.

(a) The states of an enzyme typically present during the catalysis of a substrate: the enzyme and substrate(s) (E + S) are introduced, followed by formation of the enzyme-substrate complex intermediate (ES). Next is the transition state (ES‡), followed by the enzyme-product complex intermediate (EP), and finally the enzyme and released product(s) (E + P). (b) Representative changes with time in concentration of the substrate ([S]), product ([P]), enzyme ([E]) and enzyme-substrate complex ([ES]) are shown. See Supplementary Note 1 for further discussion of important studies of methods to detect the different states of AP.

Extended Data Fig. 2 In contrast to L12, the ‘no linker’ L0 nanoantenna does not allow FAM to bind to unoccupied biotin-binding site on streptavidin nor reach the surface of bAP.

(a) Effect of free biotin binding on the nanoantenna-SA platform for the L0 (left) and L12 (right) nanoantennas. (b) Effect of free biotin binding on the nanoantenna-SA-bAP complex for the L0 and L12 nanoantennas. Discussion: (a) When the L0 or L12 nanoantennas bind to SA, their FAM fluorescence is quenched. Next, free biotin is added in excess. The fluorescence of the L0 nanoantenna does not change because its FAM moiety cannot reach any unoccupied biotin-binding sites58, thus it remains unaffected by biotin binding. In contrast, the fluorescence of the L12 nanoantenna increases because its longer length allows its FAM moiety to weakly bind at an unoccupied biotin-binding site until it is ejected by the incoming higher affinity biotin molecule55,56. (b) The L0 and L12 nanoantennas bind to SA. bAP then binds to the nanoantenna-SA platform. Next, free biotin is added in excess. The fluorescence of the L0 nanoantenna does not change upon addition of bAP because its FAM moiety cannot reach the biotin-binding sites where bAP binds and it also cannot reach the surface of bAP. Subsequent addition of free biotin also does not change its fluorescence for the same reason as without bAP and because most sites are full. In contrast, the fluorescence of the L12 nanoantenna increases upon addition of bAP because its longer length allows its FAM moiety to weakly bind at another biotin-binding site until it is ejected by the incoming high affinity biotin moiety of bAP, thereby allowing it to interact with bAP. Subsequent addition of free biotin does not substantially increase its fluorescence, indicative that the FAM moieties do not remain at the biotin-binding sites of SA. The spike observed upon addition of pNPP confirms that bAP has not been ejected. Although this experiment does not determine whether FAM is now interacting with bAP or with another location on SA, other parts of this paper demonstrate its interaction with bAP (for example, measurement of catalytic function). Conditions: 150 nM nanoantenna, 50 nM SA and 100 nM bAP in 1000 nM biotin in 200 mM Tris, 300 mM NaCl, 1 mM MgCl2, pH 7.0, 37 °C.

Extended Data Fig. 3 Molecular dynamics (MD) simulation of a possible nanoantenna-SA-bAP complex.

During the simulation, the nanoantenna locates the FAM dye closer to its binding site near the enzyme’s substrate active site. See also Video 1.

Extended Data Fig. 4 Fluorescence spikes only occur when there is a hydrolysis reaction and when the nanoantennas are close to the enzyme.

(a) Typical nanoantenna fluorescence signal used to monitor pNPP hydrolysis. (b) Addition of the reaction products, p-nitrophenol (pNP) and inorganic phosphate (Pi), does not give a fluorescence spike because there is no hydrolysis reaction. (c) Using an enzyme without phosphatase activity and which will not hydrolyze pNPP (for example, biotinylated glucose oxidase, bGOx), does not give a fluorescence spike because there is no hydrolysis reaction. (d) Here, the ‘Dummy’ nanoantenna does not have the dye (that is, no FAM) but it is still attached to SA via its biotin, while the ‘Global’ nanoantenna has FAM but it is not biotinylated and instead is free in solution. Thus, the hydrolysis reaction of pNPP still occurs, but this system does not monitor it since there is essentially no FAM-bAP interaction. (e) Here, 3′-thiolated L12 ssDNA nanoantennas were covalently attached to the lysine residues of AP (NA-AP; see Online Methods section). This nanoantenna-AP conjugate displays a spike during pNPP hydrolysis similar to the nanoantenna-SA-bAP complex. Although several synthesis steps are involved, it may be desirable for applications for which one does not wish to use the biotin-streptavidin platform. Note that the power was reduced from 635 V to 450 V due to the high baseline. (f) As a control, unattached thiolated nanoantennas and unconjugated AP do not display a spike during pNPP hydrolysis. (g) Here, a commercially prepared conjugate of SA covalently attached to AP (SA-AP) was used. The kinetic signature is shown for the PolyT L24 nanoantenna binding to SA-AP that results in fluorescence quenching, followed by pNPP hydrolysis that results in a spike. (h) Without knowledge of the SA-AP molecular weight due to an unknown number of conjugated SAs added by the manufacturer, we instead optimized using SA-AP volume (1, 2, 3, 4, 5, 7 and 14 μL SA-AP). All experiments were performed with n=1 biologically independent enzyme samples examined over 3 independent experiments near the apparent maximum (2 to 5 μL) and 1 otherwise. Data are presented as mean values ± SEM. Even after this simple optimization, however, the spike intensity during pNPP hydrolysis remains weaker compared to using the SA and bAP strategy. Overall, these results show that no matter which attachment strategy is used, and despite some being better than others, FAM will still find its binding site on the AP enzyme. Conditions: (a-d) 150 nM L12 PolyT nanoantenna, 50 nM SA, 150 nM bAP, (e,f) ~40 nM nanoantenna-bAP conjugate and (g,h) 150 nM nanoantenna, 1 to 14 μL SA-AP; 100 µM pNPP in 200 mM Tris, 300 mM NaCl, 1 mM MgCl2, pH 7.0, 37 °C.

Extended Data Fig. 5 Effect of various chemical modifiers near FAM on cDNA binding, SA binding, bAP binding, and pNPP hydrolysis.

Here, we investigated whether various chemical modifications near the dye (‘modifiers’) could affect the fluorescence signal of FAM by changing its interaction with bAP. (a) We used the L12 ssDNA FAM nanoantenna (5′ T 6-FAM) with a complementary strand containing the modifier located at the 3′-end. (b) For comparison, 1) is the cDNA without a modifier; 2) is the cDNA with phosphate; 3) is the cDNA with a hydrophobic C16 alkane chain; 4) is the cDNA with a modifier that contains a disulfide that would normally be cleaved before use to provide thiol functionality; 5) is a cDNA with the cleaved thiol. (c) Example kinetic signatures and (d) summary of all results. In short, the SA and bAP binding steps display different intensities with each modifier, but nevertheless they are qualitatively similar in all cases (that is, signal up or signal down). The exception to this is the C16 alkane chain, which results in fluorescence quenching when bAP binds. In all cases, the spike intensity during pNPP hydrolysis was similar. All experiments were performed with n=1 biologically independent enzyme samples examined over 3 independent experiments. Data are presented as mean values ± SEM. Conditions: 15 nM nanoantenna, 75 nM cDNA, 5 nM SA, 10 nM homemade bAP, 25 μM pNPP, pH 8.0, 100 mM Tris, 10 mM NaCl, 37 °C.

Extended Data Fig. 6 Effect of FAM connection and isomer on SA binding, bAP binding, and pNPP hydrolysis.

(a) In most of this study until this point, we used a L12 ssDNA nanoantenna with 5′ thymine 6-carboxyfluorescein (5′ T 6-FAM). Here, however, we also tested other FAM connections on the same DNA sequence: 5′ 6-carboxyfluorescein (5′ 6-FAM), 5′ 5-carboxyfluorescein (5′ 5-FAM) and 3′ 5-carboxyfluorescein (3′ 5-FAM). (b) Shown are the quenching of fluorescence upon SA binding, the increase of fluorescence upon bAP binding, and the transient fluorescence spike during pNPP hydrolysis. Despite the similar fluorescence emission of 5-FAM and 6-FAM when conjugated to DNA, the various FAM nanoantennas display different trends for protein binding and pNPP hydrolysis. These differences are likely due to how the chemical connection subtly affects FAM-bAP interaction. All experiments were performed with n=1 biologically independent enzyme samples examined over 3 independent experiments. Data are presented as mean values ± SEM. Conditions: 15 nM nanoantenna, 5 nM SA, 10 nM homemade bAP, 30 μM pNPP, pH 8.0, 100 mM Tris, 10 mM NaCl, 30 °C. PMT voltage = 800 V.

Extended Data Fig. 7 Molecular dynamics (MD) trajectories of dyes and/or substrate on AP.

The 100 ns MD simulation of AP with or without a dye (FAM, CAL, Cy3) and with or without the substrate pNPP. We selected the lowest energy pose (see Supplementary Fig. 13) and the next lowest energy pose that does not overlap with it and that has the linker location exposed. Dyes and pNPP are circled for visual clarity. In all cases, we observed that pNPP remains bound at the active sites. For AP with FAM and with or without pNPP, the position of FAM remains unchanged. For AP with CAL and with or without pNPP, the CAL dye does not have a stable position in either case. For AP-Cy3 without pNPP, the Cy3 dye position does not change, but with pNPP bound, the dye position can change, and it even dissociates from the surface. We emphasize that these MD simulations represent a possible signaling mechanism (see main text), and not a definite proposal.

Extended Data Fig. 8 Excitation and emission spectra of double-dye dsDNA nanoantenna (FAM-CAL) suggest dye stacking.

The excitation spectra (dashed line) and emission spectra (solid line) of the formation of the double-dye dsDNA nanoantenna: starting with ssDNA, after binding of cDNA, after binding of SA, and after binding of bAP. In (a), only the FAM dye is present, and excitation and emission of FAM wavelengths (λem = 520 nm and λex = 498 nm) are relatively unaffected by the addition of the complementary DNA (of note, the dsDNA antenna displays little sensitivity to SA and bAP attachment relative to ssDNA nanoantennas, see Fig. 1). In (b), only the CAL dye is present, and excitation and emission of CAL wavelengths (λem = 561 nm and λex = 540 nm) are relatively unaffected by the addition of the complementary DNA. In (c) and (d), both dyes were present after the cDNA step (that is, the systems were chemically identical), but (c) was measured with the FAM wavelengths and (d) with the CAL wavelengths. We observed that both the FAM and CAL excitation and emission spectra are drastically affected when proximal to the other dye. This remains true even following the addition of the SA and bAP proteins. This decrease of signal intensity is likely attributable to a contact-mediated quenching mechanism between the dyes65. These dyes seem to remain stacked even after the nanoantenna has moved to SA and bAP.

Extended Data Fig. 9 Comparison of methods and error.

(a) Classic Michaelis-Menten method to determined kcat and KM values for the 4MUP substrate using the initial rates of 4MU product generation. (b) Nanoantenna ‘one-shot’ method to determined kcat and KM values using the FAM fluorescence spike obtained during 4MUP hydrolysis. The values determined using both methods displayed good agreement. This analysis was performed several months after our values obtained for 4MUP in Fig. 3, which also shows the reproducibility of the method. (c) We further compared the accuracy of the ‘one-shot’ nanoantenna method over a similar approach performed using the 4MU progress curve under the same conditions. We found that a ‘one shot’ 4MU progress curve over-estimated both the kcat and KM of the enzyme-substrate system. See relevant literature for fitting of progress curves. Conditions were 100 mM Tris, 10 mM NaCl, pH 8.0, 37 °C; also 150 nM nanoantenna, 50 nM SA, 20 nM bAP, and 300 μM 4MUP in (b and c), and the same buffer and temperature but 37.5 nM nanoantenna, 12.5 nM SA, 5 nM bAP, and 2 μM, 5 μM, 10 μM, 20 μM, 40 μM, 80 μM, 140 μM or 350 μM 4MUP in (a). In the latter two, the concentration of bAP was reduced to facilitate measurement of low 4MUP concentrations, and the nanoantenna and SA concentrations were reduced proportionately. All experiments were performed with n=1 biologically independent enzyme samples examined over 3 independent experiments. (24 measurements for the classic Michaelis-Menten via initial rates and 3 each for the nanoantenna spike and product progress curve). Data are presented as mean values ± SEM.

Extended Data Fig. 10 Theoretical nanoantenna kinetic signatures for different types of inhibitors.

Here, we generated the expected spike profile of a theoretical system with the parameters: kcat = 100 s−1, KM = 10 µM, [enzyme] = 100 nM, [substrate] = 1000 µM, and [inhibitor] = 125 µM. Shown are the effects of (a) competitive inhibitors with Ki = 100 µM and Ki = 1 µM, (b) uncompetitive inhibitor with Ki = 100 µM, and (c) non-competitive inhibitor with Ki = 100 µM.

Supplementary information

Supplementary Information

Supplementary Figs. 1–35, Tables 1–4 and Notes 1 and 2. List of reagents. List of oligonucleotide sequences. Script for fitting kinetic data in MATLAB. Supplementary refs.

Reporting Summary

Supplementary Video 1

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Harroun, S.G., Lauzon, D., Ebert, M.C.C.J.C. et al. Monitoring protein conformational changes using fluorescent nanoantennas. Nat Methods 19, 71–80 (2022). https://doi.org/10.1038/s41592-021-01355-5

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue date:

  • DOI: https://doi.org/10.1038/s41592-021-01355-5

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing