Single-molecule digital sizing of proteins in solution.
Journal
Nature communications
ISSN: 2041-1723
Titre abrégé: Nat Commun
Pays: England
ID NLM: 101528555
Informations de publication
Date de publication:
04 Sep 2024
04 Sep 2024
Historique:
received:
14
07
2023
accepted:
23
07
2024
medline:
5
9
2024
pubmed:
5
9
2024
entrez:
4
9
2024
Statut:
epublish
Résumé
The physical characterization of proteins in terms of their sizes, interactions, and assembly states is key to understanding their biological function and dysfunction. However, this has remained a difficult task because proteins are often highly polydisperse and present as multicomponent mixtures. Here, we address this challenge by introducing single-molecule microfluidic diffusional sizing (smMDS). This approach measures the hydrodynamic radius of single proteins and protein assemblies in microchannels using single-molecule fluorescence detection. smMDS allows for ultrasensitive sizing of proteins down to femtomolar concentrations and enables affinity profiling of protein interactions at the single-molecule level. We show that smMDS is effective in resolving the assembly states of protein oligomers and in characterizing the size of protein species within complex mixtures, including fibrillar protein aggregates and nanoscale condensate clusters. Overall, smMDS is a highly sensitive method for the analysis of proteins in solution, with wide-ranging applications in drug discovery, diagnostics, and nanobiotechnology.
Identifiants
pubmed: 39231922
doi: 10.1038/s41467-024-50825-9
pii: 10.1038/s41467-024-50825-9
doi:
Substances chimiques
Proteins
0
Solutions
0
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
7740Informations de copyright
© 2024. The Author(s).
Références
Goodsell, D. S. The Machinery of Life. The Machinery of Life (Springer, New York, NY, 2009). https://doi.org/10.1007/978-0-387-84925-6 .
Neet, K. E. & Lee, J. C. Biophysical characterization of proteins in the post-genomic era of proteomics. Mol. Cell Proteom. 1, 415–420 (2002).
doi: 10.1074/mcp.R200003-MCP200
Knowles, T. P. J., Vendruscolo, M. & Dobson, C. M. The amyloid state and its association with protein misfolding diseases. Nat. Rev. Mol. Cell Biol. 15, 384–396 (2014).
pubmed: 24854788
doi: 10.1038/nrm3810
Eliezer, D. Biophysical characterization of intrinsically disordered proteins. Curr. Opin. Struct. Biol. 19, 23–30 (2009).
pubmed: 19162471
pmcid: 2728036
doi: 10.1016/j.sbi.2008.12.004
Bemporad, F. & Chiti, F. Protein misfolded oligomers: experimental approaches, mechanism of formation, and structure-toxicity relationships. Chem. Biol. 19, 315–327 (2012).
pubmed: 22444587
doi: 10.1016/j.chembiol.2012.02.003
Ha, J., Park, H., Park, J. & Park, S. B. Recent advances in identifying protein targets in drug discovery. Cell Chem. Biol. 28, 394–423 (2021).
pubmed: 33357463
doi: 10.1016/j.chembiol.2020.12.001
Rapposelli, S., Gaudio, E., Bertozzi, F. & Gul, S. Editorial: Protein–protein interactions: drug discovery for the future. Front Chem. 9, 1049 (2021).
doi: 10.3389/fchem.2021.811190
Shen, Y. et al. From protein building blocks to functional materials. ACS Nano 15, 5819–5837 (2021).
pubmed: 33760579
pmcid: 8155333
doi: 10.1021/acsnano.0c08510
Houde, D. J. & Berkowitz, S. A. Biophysical Characterization of Proteins in Developing Biopharmaceuticals. Biophysical Characterization of Proteins in Developing Biopharmaceuticals (Elsevier, 2020). https://doi.org/10.1016/C2017-0-03008-2 .
Renaud, J. P. et al. Biophysics in drug discovery: impact, challenges and opportunities. Nat. Rev. Drug Discov. 15, 679–698 (2016).
Hamley, I. W. Protein Assemblies: Nature-Inspired and Designed Nanostructures. Biomacromolecules 20, 1829–1848 (2019).
pubmed: 30912925
pmcid: 7007009
doi: 10.1021/acs.biomac.9b00228
Modena, M. M., Rühle, B., Burg, T. P. & Wuttke, S. Nanoparticle characterization: What to measure? Adv. Mater. 31, 1901556 (2019).
doi: 10.1002/adma.201901556
Lin, P. C., Lin, S., Wang, P. C. & Sridhar, R. Techniques for physicochemical characterization of nanomaterials. Biotechnol. Adv. 32, 711 (2014).
pubmed: 24252561
doi: 10.1016/j.biotechadv.2013.11.006
Walport, L. J., Low, J. K. K., Matthews, J. M. & MacKay, J. P. The characterization of protein interactions – what, how and how much? Chem. Soc. Rev. 50, 12292–12307 (2021).
pubmed: 34581717
doi: 10.1039/D1CS00548K
Tamara, S., Den Boer, M. A. & Heck, A. J. R. High-resolution native mass spectrometry. Chem. Rev. 122, 7269–7326 (2022).
pubmed: 34415162
doi: 10.1021/acs.chemrev.1c00212
Barth, H. G., Jackson, C. & Boyes, B. E. Size exclusion Chromatography. Anal. Chem. 66, 595–620 (1994).
doi: 10.1021/ac00084a022
Nguyen, H. H., Park, J., Kang, S. & Kim, M. Surface plasmon resonance: a versatile technique for biosensor applications. Sensors 15, 10481–10510 (2015).
pubmed: 25951336
pmcid: 4481982
doi: 10.3390/s150510481
Fekete, S., Guillarme, D., Sandra, P. & Sandra, K. Chromatographic, electrophoretic, and mass spectrometric methods for the analytical characterization of protein biopharmaceuticals. Anal. Chem. 88, 480–507 (2016).
pubmed: 26629607
doi: 10.1021/acs.analchem.5b04561
Velázquez Campoy, A. & Freire, E. ITC in the post-genomic era…? Priceless. Biophys. Chem. 115, 115–124 (2005).
pubmed: 15752592
doi: 10.1016/j.bpc.2004.12.015
Cole, J. L., Lary, J. W., P. Moody, T. & Laue, T. M. Analytical ultracentrifugation: sedimentation velocity and sedimentation equilibrium. Methods Cell Biol. 84, 143 (2008).
pubmed: 17964931
pmcid: 2711687
doi: 10.1016/S0091-679X(07)84006-4
Stetefeld, J., McKenna, S. A. & Patel, T. R. Dynamic light scattering: a practical guide and applications in biomedical sciences. Biophys. Rev. 8, 409 (2016).
pubmed: 28510011
pmcid: 5425802
doi: 10.1007/s12551-016-0218-6
Haustein, E. & Schwille, P. Fluorescence correlation Spectroscopy: Novel variations of an established technique. Annu Rev. Biophys. Biomol. Struct. 36, 151–169 (2007).
pubmed: 17477838
doi: 10.1146/annurev.biophys.36.040306.132612
Yu, L. et al. A comprehensive review of fluorescence correlation spectroscopy. Front Phys. 9, 644450 (2021).
doi: 10.3389/fphy.2021.644450
Puchalla, J., Krantz, K., Austin, R. & Rye, H. Burst analysis spectroscopy: A versatile single-particle approach for studying distributions of protein aggregates and fluorescent assemblies. Proc. Natl Acad. Sci. USA 105, 14400–14405 (2008).
pubmed: 18780782
pmcid: 2567176
doi: 10.1073/pnas.0805969105
Gambin, Y. et al. Confocal spectroscopy to study dimerization, oligomerization and aggregation of proteins: a practical guide. Int. J. Mol. Sci. 2016 17, 655 (2016).
doi: 10.3390/ijms17050655
Dresser, L. et al. Amyloid-β oligomerization monitored by single-molecule stepwise photobleaching. Methods 193, 80–95 (2021).
pubmed: 32544592
pmcid: 8336786
doi: 10.1016/j.ymeth.2020.06.007
Arant, R. J. & Ulbrich, M. H. Deciphering the subunit composition of multimeric proteins by counting photobleaching steps. Chemphyschem. 15, 600–605 (2014).
pubmed: 24481650
doi: 10.1002/cphc.201301092
Sengupta, P., Garai, K., Balaji, J., Periasamy, N. & Maiti, S. Measuring size distribution in highly heterogeneous systems with fluorescence correlation spectroscopy. Biophys. J. 84, 1977 (2003).
pubmed: 12609900
pmcid: 1302767
doi: 10.1016/S0006-3495(03)75006-1
Hillger, F., Nettels, D., Dorsch, S. & Schuler, B. Detection and analysis of protein aggregation with confocal single molecule fluorescence spectroscopy. J. Fluoresc. 17, 759–765 (2007).
pubmed: 17447125
doi: 10.1007/s10895-007-0187-z
Mendes, A., Heil, H. S., Coelho, S., Leterrier, C. & Henriques, R. Mapping molecular complexes with super-resolution microscopy and single-particle analysis. Open Biol. 12, (2022).
Sydor, A. M., Czymmek, K. J., Puchner, E. M. & Mennella, V. Super-resolution microscopy: from single molecules to supramolecular assemblies. Trends Cell Biol. 25, 730–748 (2015).
pubmed: 26546293
doi: 10.1016/j.tcb.2015.10.004
Young, G. et al. Quantitative mass imaging of single biological macromolecules. Science 360, 423–427 (2018).
pubmed: 29700264
pmcid: 6103225
doi: 10.1126/science.aar5839
Piliarik, M. & Sandoghdar, V. Direct optical sensing of single unlabelled proteins and super-resolution imaging of their binding sites. Nat. Commun. 5, 1–8 (2014).
doi: 10.1038/ncomms5495
Taylor, R. W. & Sandoghdar, V. Interferometric Scattering microscopy: seeing single nanoparticles and molecules via Rayleigh scattering. Nano Lett. 19, 4827–4835 (2019).
pubmed: 31314539
pmcid: 6750867
doi: 10.1021/acs.nanolett.9b01822
Young, G. & Kukura, P. Interferometric Scattering Microscopy. 10.1146/annurev-physchem-050317-021247 70, 301–322 (2019).
Liebthal, M., Kushwah, M. S., Kukura, P. & Dietz, K. J. Single molecule mass photometry reveals the dynamic oligomerization of human and plant peroxiredoxins. iScience 24, (2021).
Fineberg, A., Surrey, T. & Kukura, P. Quantifying the Monomer-Dimer equilibrium of Tubulin with mass photometry. J. Mol. Biol. 432, 6168–6172 (2020).
pubmed: 33068635
pmcid: 7763485
doi: 10.1016/j.jmb.2020.10.013
Sonn-Segev, A. et al. Quantifying the heterogeneity of macromolecular machines by mass photometry. Nat. Commun.11, 1–10 (2020).
Arter, W. E., Levin, A., Krainer, G. & Knowles, T. P. J. Microfluidic approaches for the analysis of protein–protein interactions in solution. Biophys. Rev. 1–11 https://doi.org/10.1007/s12551-020-00679-4 (2020).
Charmet, J., Arosio, P. & Knowles, T. P. J. Microfluidics for protein biophysics. J. Mol. Biol. 430, 565–580 (2018).
pubmed: 29289566
doi: 10.1016/j.jmb.2017.12.015
Otzen, D. E., Buell, A. K. & Jensen, H. Microfluidics and the quantification of biomolecular interactions. Curr. Opin. Struct. Biol. 70, 8–15 (2021).
pubmed: 33831785
doi: 10.1016/j.sbi.2021.02.006
Arosio, P. et al. Microfluidic diffusion analysis of the sizes and interactions of proteins under native solution conditions. ACS Nano 10, 333–341 (2016).
pubmed: 26678709
doi: 10.1021/acsnano.5b04713
Zhang, Y. et al. On-chip measurements of protein unfolding from direct observations of micron-scale diffusion. Chem. Sci. 9, 3503–3507 (2018).
pubmed: 29780480
pmcid: 5934698
doi: 10.1039/C7SC04331G
Challa, P. K. et al. Real-time intrinsic fluorescence visualization and sizing of proteins and protein complexes in microfluidic devices. Anal. Chem. 90, 3849–3855 (2018).
pubmed: 29451779
doi: 10.1021/acs.analchem.7b04523
Wright, M. A. et al. Cooperative assembly of Hsp70 subdomain clusters. Biochemistry 57, 3641–3649 (2018).
pubmed: 29763298
doi: 10.1021/acs.biochem.8b00151
Fiedler, S. et al. Antibody affinity governs the inhibition of SARS-CoV-2 Spike/ACE2 binding in patient serum. ACS Infect. Dis. 7, 2362–2369 (2021).
pubmed: 33876632
doi: 10.1021/acsinfecdis.1c00047
Schneider, M. M. et al. The Hsc70 disaggregation machinery removes monomer units directly from α-synuclein fibril ends. Nat. Commun. 12, 1–11 (2021).
Linse, S. et al. Kinetic fingerprints differentiate the mechanisms of action of anti-Aβ antibodies. Nat. Struct. Mol. Biol. 27, 1125–1133 (2020).
pubmed: 32989305
doi: 10.1038/s41594-020-0505-6
Baron, J. et al. FULL-MDS: Fluorescent universal lipid labeling for microfluidic diffusional sizing. Anal. Chem. 95, 587–593 (2023).
pubmed: 36574263
Fiedler, S. et al. Serological fingerprints link antiviral activity of therapeutic antibodies to affinity and concentration. Sci. Rep. 12, 1–10 (2022).
doi: 10.1038/s41598-022-22214-z
Scheidt, T. et al. Secondary nucleation and elongation occur at different sites on Alzheimer’s amyloid-b aggregates. Sci. Adv. 5, eaau3112 (2019).
pubmed: 31001578
pmcid: 6469941
doi: 10.1126/sciadv.aau3112
Yates, E. V. et al. Latent analysis of unmodified biomolecules and their complexes in solution with attomole detection sensitivity. Nat. Chem. 7, 802–809 (2015).
pubmed: 26391079
doi: 10.1038/nchem.2344
Schneider, M. M. et al. Microfluidic antibody affinity profiling of alloantibody-HLA interactions in human serum. Biosens. Bioelectron. 228, 115196 (2023).
pubmed: 36921387
doi: 10.1016/j.bios.2023.115196
Schneider, M. M. et al. Microfluidic characterisation reveals broad range of SARS-CoV-2 antibody affinity in human plasma. Life Sci. Alliance 5, e202101270 (2022).
pubmed: 34848436
doi: 10.26508/lsa.202101270
Moser, M. R. & Baker, C. A. Taylor dispersion analysis in fused silica capillaries: a tutorial review. Anal. Methods 13, 2357–2373 (2021).
pubmed: 33999088
doi: 10.1039/D1AY00588J
Poulsen, N. N. et al. Flow induced dispersion analysis rapidly quantifies proteins in human plasma samples. Analyst 140, 4365–4369 (2015).
pubmed: 26031223
doi: 10.1039/C5AN00697J
Single Molecule Detection in Solution. Single Molecule Detection in Solution https://doi.org/10.1002/3527600809 (2002).
Shashkova, S. & Leake, M. C. Single-molecule fluorescence microscopy review: shedding new light on old problems. Biosci. Rep. 37, 20170031 (2017).
doi: 10.1042/BSR20170031
Nie, S., Chiu, D. T. & Zare, R. N. Real-time detection of single molecules in solution by confocal fluorescence microscopy. Anal. Chem. 67, 2849–2857 (1995).
doi: 10.1021/ac00113a019
Krainer, G. et al. Direct digital sensing of protein biomarkers in solution. Nat. Commun. 14, 1–21 (2023).
doi: 10.1038/s41467-023-35792-x
Horrocks, M. H. et al. Single molecule fluorescence under conditions of fast flow. Anal. Chem. 84, 179–185 (2012).
pubmed: 22147688
doi: 10.1021/ac202313d
Zijlstra, N. et al. Rapid Microfluidic Dilution for Single-Molecule Spectroscopy of Low-Affinity Biomolecular Complexes. Angew. Chem. Int. Ed. 56, 7126–7129 (2017).
doi: 10.1002/anie.201702439
Tyagi, S. et al. Continuous throughput and long-term observation of single-molecule FRET without immobilization. Nat. Methods 11, 297–300 (2014).
pubmed: 24441935
doi: 10.1038/nmeth.2809
Kim, S. et al. High-throughput single-molecule optofluidic analysis. Nat. Methods 8, 242–245 (2011).
pubmed: 21297618
pmcid: 3075913
doi: 10.1038/nmeth.1569
Paganini, C. et al. Rapid characterization and quantification of extracellular vesicles by fluorescence-based microfluidic diffusion sizing. Adv. Health. Mater. 11, 2100021 (2022).
doi: 10.1002/adhm.202100021
Scheidt, T. et al. Multidimensional protein characterisation using microfluidic post-column analysis. Lab Chip 20, 2663–2673 (2020).
pubmed: 32588855
doi: 10.1039/D0LC00219D
Armstrong, J. K., Wenby, R. B., Meiselman, H. J. & Fisher, T. C. The hydrodynamic radii of macromolecules and their effect on red blood cell aggregation. Biophys. J. 87, 4259–4270 (2004).
pubmed: 15361408
pmcid: 1304934
doi: 10.1529/biophysj.104.047746
Jiang, X. et al. Quantitative analysis of the protein corona on FePt nanoparticles formed by transferrin binding. J. R. Soc. Interface 7 (2010).
Akbarzadehlaleh, P., Mirzaei, M., Mashahdi-Keshtiban, M., Shamsasenjan, K. & Heydari, H. PEGylated human serum Albumin: Review of PEGylation, purification and characterization methods. Adv. Pharm. Bull. 6, 309–317 (2016).
pubmed: 27766215
pmcid: 5071794
doi: 10.15171/apb.2016.043
Dynamic Biosensors. List of protein hydrodynamic diameters DH. https://www.dynamic-biosensors.com/project/list-of (2022).
Andersson, L. O., Borg, H. & Mikaelsson, M. Molecular weight estimations of proteins by electrophoresis in polyacrylamide gels of graded porosity. FEBS Lett. 20, 199–202 (1972).
pubmed: 11946415
doi: 10.1016/0014-5793(72)80793-2
Vendelin, M. & Birkedal, R. Anisotropic diffusion of fluorescently labeled ATP in rat cardiomyocytes determined by raster image correlation spectroscopy. Am. J. Physiol. Cell Physiol. 295, C1302–C1315 (2008).
pubmed: 18815224
pmcid: 2584976
doi: 10.1152/ajpcell.00313.2008
Study of Protein Hydrodynamics with Light Scattering. https://www.brookhaveninstruments.com/study-of-protein-hydrodynamics-with-light-scattering-size-and-charge-of-lysozyme/ .
Size Measurement of the Proteins RNAse A and Proteinase K. https://www.brookhaveninstruments.com/size-measurement-of-the-proteins-rnase-a-and-proteinase-k-and-the-effects-of-protein-binding/ .
Falke, M. et al. α-Synuclein-derived lipoparticles in the study of α-Synuclein amyloid fibril formation. Chem. Phys. Lipids 220, 57–65 (2019).
pubmed: 30826264
pmcid: 6451039
doi: 10.1016/j.chemphyslip.2019.02.009
Schneider, S. Determination of Protein Molecular Weight and Size Using the Agilent 1260 Infinity Multi-Detector Bio-SEC Solution with Advanced Light Scattering Detection. Appl. Note 1–6 (2014).
Arter, W. E. et al. Rapid structural, kinetic, and immunochemical analysis of Alpha-Synuclein Oligomers in solution. Nano Lett. 20, 8163–8169 (2020).
pubmed: 33079553
pmcid: 7116857
doi: 10.1021/acs.nanolett.0c03260
Lakayan, D., Haselberg, R., Gahoual, R., Somsen, G. W. & Kool, J. Affinity profiling of monoclonal antibody and antibody-drug-conjugate preparations by coupled liquid chromatography-surface plasmon resonance biosensing. Anal. Bioanal. Chem. 410, 7837–7848 (2018).
pubmed: 30328504
pmcid: 6244757
doi: 10.1007/s00216-018-1414-y
Biddison, W. E. et al. Structural studies of an HLA-A03 alloantigenic epitope defined by a human hybridoma antibody. Immunogenetics 30, 54–57 (1989).
pubmed: 2473031
doi: 10.1007/BF02421471
Ivanyi, D. & van de Meugheuvel, W. A monomorphic HLA-specific monoclonal antibody, W6/32, reacts with the H-2Db molecule of normal mouse lymphocytes. Immunogenetics 20, 699–703 (1984).
pubmed: 6510995
doi: 10.1007/BF00430328
Aprile, F. A. et al. Rational design of a conformation-specific antibody for the quantification of Aβ oligomers. Proc. Natl Acad. Sci. USA 117, 13509–13518 (2020).
pubmed: 32493749
pmcid: 7306997
doi: 10.1073/pnas.1919464117
Levy, E. D. & Teichmann, S. Structural, evolutionary, and assembly principles of protein oligomerization. Prog. Mol. Biol. Transl. Sci. 117, 25–51 (2013).
pubmed: 23663964
doi: 10.1016/B978-0-12-386931-9.00002-7
Ali, M. H. & Imperiali, B. Protein oligomerization: how and why. Bioorg. Med. Chem. 13, 5013–5020 (2005).
pubmed: 15993087
doi: 10.1016/j.bmc.2005.05.037
Gell, D. A., Grant, R. P. & MacKay, J. P. The detection and quantitation of protein oligomerization. Adv. Exp. Med. Biol. 747, 19–41 (2012).
pubmed: 22949109
doi: 10.1007/978-1-4614-3229-6_2
Kulenkampff, K., Wolf Perez, A. M., Sormanni, P., Habchi, J. & Vendruscolo, M. Quantifying misfolded protein oligomers as drug targets and biomarkers in Alzheimer and Parkinson diseases. Nat. Rev. Chem. 5, 277–294 (2021).
pubmed: 37117282
doi: 10.1038/s41570-021-00254-9
Kumari, N. & Yadav, S. Modulation of protein oligomerization: An overview. Prog. Biophys. Mol. Biol. 149, 99–113 (2019).
pubmed: 30872157
doi: 10.1016/j.pbiomolbio.2019.03.003
Griffin, M. D. W. & Gerrard, J. A. The relationship between oligomeric state and protein function. Adv. Exp. Med Biol. 747, 74–90 (2012).
pubmed: 22949112
doi: 10.1007/978-1-4614-3229-6_5
Du, X. Y., Xie, X. X. & Liu, R. T. The role of α-Synuclein Oligomers in Parkinson’s disease. Int. J. Mol. Sci. 21, 8645 (2020).
pubmed: 33212758
pmcid: 7697105
doi: 10.3390/ijms21228645
Musteikytė, G. et al. Interactions of α-synuclein oligomers with lipid membranes. Biochim. Biophys. Acta (BBA) - Biomembr. 1863, 183536 (2020).
doi: 10.1016/j.bbamem.2020.183536
Alam, P., Bousset, L., Melki, R. & Otzen, D. E. α-synuclein oligomers and fibrils: a spectrum of species, a spectrum of toxicities. J. Neurochem. 150, 522–534 (2019).
pubmed: 31254394
doi: 10.1111/jnc.14808
Chen, S. W. et al. Structural characterization of toxic oligomers that are kinetically trapped during α-synuclein fibril formation. Proc. Natl Acad. Sci. USA 112, E1994–E2003 (2015).
pubmed: 25855634
pmcid: 4413268
Lorenzen, N. et al. The role of stable α-synuclein oligomers in the molecular events underlying amyloid formation. J. Am. Chem. Soc. 136, 3859–3868 (2014).
pubmed: 24527756
doi: 10.1021/ja411577t
Fusco, G. et al. Structural basis of membrane disruption and cellular toxicity by α-synuclein oligomers. Science 358, 1440–1443 (2017).
pubmed: 29242346
doi: 10.1126/science.aan6160
Lee, J. E. et al. Mapping surface hydrophobicity of α-synuclein oligomers at the nanoscale. Nano Lett. 18, 7494–7501 (2018).
pubmed: 30380895
pmcid: 6295917
doi: 10.1021/acs.nanolett.8b02916
Staats, R. et al. Screening of small molecules using the inhibition of oligomer formation in α-synuclein aggregation as a selection parameter. Commun. Chem. 3, 1–9 (2020).
doi: 10.1038/s42004-020-00412-y
Apetri, M. M., Maiti, N. C., Zagorski, M. G., Carey, P. R. & Anderson, V. E. Secondary structure of α-Synuclein Oligomers: Characterization by Raman and atomic force microscopy. J. Mol. Biol. 355, 63–71 (2006).
pubmed: 16303137
doi: 10.1016/j.jmb.2005.10.071
Lowe, R., Pountney, D. L., Jensen, P. H., Gai, W. P. & Voelcker, N. H. Calcium(II) selectively induces α-synuclein annular oligomers via interaction with the C-terminal domain. Protein Sci. 13, 3245–3252 (2004).
pubmed: 15537754
pmcid: 2287302
doi: 10.1110/ps.04879704
Norris, E. H. et al. Reversible inhibition of α-synuclein fibrillization by dopaminochrome-mediated conformational alterations. J. Biol. Chem. 280, 21212–21219 (2005).
pubmed: 15817478
doi: 10.1074/jbc.M412621200
Paslawski, W. et al. High stability and cooperative unfolding of α-Synuclein Oligomers. Biochemistry 53, 6252–6263 (2014).
pubmed: 25216651
doi: 10.1021/bi5007833
Wright, J. A., Wang, X. & Brown, D. R. Unique copper-induced oligomers mediate alpha-synuclein toxicity. FASEB J. 23, 2384–2393 (2009).
pubmed: 19325037
doi: 10.1096/fj.09-130039
Hayden, E. Y. et al. Heme stabilization of α-Synuclein oligomers during amyloid fibril formation. Biochemistry 54, 4599–4610 (2015).
pubmed: 26161848
doi: 10.1021/acs.biochem.5b00280
Pieri, L., Madiona, K., Bousset, L. & Melki, R. Fibrillar α-synuclein and huntingtin exon 1 assemblies are toxic to the cells. Biophys. J. 102, 2894–2905 (2012).
pubmed: 22735540
pmcid: 3379023
doi: 10.1016/j.bpj.2012.04.050
Kaufmann, T. J., Harrison, P. M., Richardson, M. J. E., Pinheiro, T. J. T. & Wall, M. J. Intracellular soluble α-synuclein oligomers reduce pyramidal cell excitability. J. Physiol. 594, 2751–2772 (2016).
pubmed: 26915902
pmcid: 4865569
doi: 10.1113/JP271968
Qin, Z. et al. Effect of 4-hydroxy-2-nonenal modification on α-synuclein aggregation. J. Biol. Chem. 282, 5862–5870 (2007).
pubmed: 17189262
doi: 10.1074/jbc.M608126200
Stefanovic, A. N. D., Lindhoud, S., Semerdzhiev, S. A., Claessens, M. M. A. E. & Subramaniam, V. Oligomers of Parkinson’s disease-related α-synuclein mutants have similar structures but distinctive membrane permeabilization properties. Biochemistry 54, 3142–3150 (2015).
pubmed: 25909158
doi: 10.1021/bi501369k
Giehm, L., Svergun, D. I., Otzen, D. E. & Vestergaard, B. Low-resolution structure of a vesicle disrupting α-synuclein oligomer that accumulates during fibrillation. Proc. Natl Acad. Sci. USA 108, 3246–3251 (2011).
pubmed: 21300904
pmcid: 3044375
doi: 10.1073/pnas.1013225108
Fecchio, C. et al. α-Synuclein Oligomers induced by Docosahexaenoic acid affect membrane integrity. PLoS One 8, e82732 (2013).
pubmed: 24312431
pmcid: 3843715
doi: 10.1371/journal.pone.0082732
Ross, C. A. & Poirier, M. A. Protein aggregation and neurodegenerative disease. Nat. Med. 10, S10–S17 (2004).
pubmed: 15272267
doi: 10.1038/nm1066
Lashuel, H. A. Do Lewy bodies contain alpha-synuclein fibrils? and Does it matter? A brief history and critical analysis of recent reports. Neurobiol. Dis. 141, 104876 (2020).
pubmed: 32339655
doi: 10.1016/j.nbd.2020.104876
Meisl, G. et al. Differences in nucleation behavior underlie the contrasting aggregation kinetics of the Aβ40 and Aβ42 peptides. Proc. Natl Acad. Sci. 111, 9384–9389 (2014).
pubmed: 24938782
pmcid: 4084462
doi: 10.1073/pnas.1401564111
Shin, Y. & Brangwynne, C. P. Liquid phase condensation in cell physiology and disease. Science 357, eaaf4382 (2017).
pubmed: 28935776
doi: 10.1126/science.aaf4382
Alberti, S. & Dormann, D. Liquid–liquid phase separation in disease. Annu Rev. Genet. 53, 171–194 (2019).
pubmed: 31430179
doi: 10.1146/annurev-genet-112618-043527
Brangwynne, C. P., Tompa, P. & Pappu, R. V. Polymer physics of intracellular phase transitions. Nat. Phys. 11, 899–904 (2015).
doi: 10.1038/nphys3532
Boeynaems, S. et al. Protein phase separation: a new phase in cell biology. Trends Cell Biol. 28, 420–435 (2018).
pubmed: 29602697
pmcid: 6034118
doi: 10.1016/j.tcb.2018.02.004
Krainer, G. et al. Reentrant liquid condensate phase of proteins is stabilized by hydrophobic and non-ionic interactions. Nat. Commun. 12, 1–14 (2021).
doi: 10.1038/s41467-021-21181-9
Maharana, S. et al. RNA buffers the phase separation behavior of prion-like RNA binding proteins. Science 360, 918–921 (2018).
pubmed: 29650702
pmcid: 6091854
doi: 10.1126/science.aar7366
Patel, A. et al. A liquid-to-solid phase transition of the ALS Protein FUS accelerated by disease mutation. Cell 162, 1066–1077 (2015).
pubmed: 26317470
doi: 10.1016/j.cell.2015.07.047
Kar, M. et al. Phase-separating RNA-binding proteins form heterogeneous distributions of clusters in subsaturated solutions. Proc. Natl Acad. Sci. USA 119, e2202222119 (2022).
pubmed: 35787038
pmcid: 9282234
doi: 10.1073/pnas.2202222119
Wen, J. et al. Conformational expansion of Tau in condensates promotes irreversible aggregation. J. Am. Chem. Soc. 143, 13056–13064 (2021).
pubmed: 34374536
doi: 10.1021/jacs.1c03078
Li, P. et al. Phase transitions in the assembly of multivalent signalling proteins. Nature 483, 336–340 (2012).
pubmed: 22398450
pmcid: 3343696
doi: 10.1038/nature10879
Seim, I. et al. Dilute phase oligomerization can oppose phase separation and modulate material properties of a ribonucleoprotein condensate. Proc. Natl Acad. Sci. USA 119, (2022).
Fleming, P. J. & Fleming, K. G. HullRad: Fast calculations of folded and disordered protein and nucleic acid hydrodynamic properties. Biophys. J. 114, 856–869 (2018).
pubmed: 29490246
pmcid: 5984988
doi: 10.1016/j.bpj.2018.01.002
Burke, K. A., Janke, A. M., Rhine, C. L. & Fawzi, N. L. Residue-by-residue view of in vitro FUS granules that bind the C-terminal domain of RNA Polymerase II. Mol. Cell 60, 231–241 (2015).
pubmed: 26455390
pmcid: 4609301
doi: 10.1016/j.molcel.2015.09.006
Murthy, A. C. et al. Molecular interactions underlying liquid−liquid phase separation of the FUS low-complexity domain. Nat. Struct. Mol. Biol. 26, 637–648 (2019).
pubmed: 31270472
pmcid: 6613800
doi: 10.1038/s41594-019-0250-x
Ahlers, J. et al. The key role of solvent in condensation: Mapping water in liquid-liquid phase-separated FUS. Biophys. J. 120, 1266–1275 (2021).
pubmed: 33515602
pmcid: 8059208
doi: 10.1016/j.bpj.2021.01.019
Li, Q., Babinchak, W. M. & Surewicz, W. K. Cryo-EM structure of amyloid fibrils formed by the entire low complexity domain of TDP-43. Nat. Commun. 12, 1–8 (2021).
Cisse, I. I. et al. Real-time dynamics of RNA polymerase II clustering in live human cells. Science 341, 664–667 (2013).
pubmed: 23828889
doi: 10.1126/science.1239053
Pancholi, A. et al. RNA polymerase II clusters form in line with surface condensation on regulatory chromatin. Mol. Syst. Biol. 17, (2021).
Longfield, S. F. et al. Tau forms synaptic nano-biomolecular condensates controlling the dynamic clustering of recycling synaptic vesicles. Nat. Commun. 14, 1–20 (2023).
Ray, S. et al. Mass photometric detection and quantification of nanoscale α-synuclein phase separation. Nat. Chem. 15, 1306–1316 (2023).
pubmed: 37337111
doi: 10.1038/s41557-023-01244-8
Hohlbein, J., Craggs, T. D. & Cordes, T. Alternating-laser excitation: single-molecule FRET and beyond. Chem. Soc. Rev. 43, 1156–1171 (2014).
pubmed: 24037326
doi: 10.1039/C3CS60233H
Schuler, B. Single-molecule FRET of protein structure and dynamics - a primer. J. Nanobiotechnology 11, S2 (2013).
pubmed: 24565277
pmcid: 4029180
doi: 10.1186/1477-3155-11-S1-S2
Orte, A., Clarke, R. & Klenerman, D. Single-molecule two-colour coincidence detection to probe biomolecular associations. Biochem Soc. Trans. 38, 914–918 (2010).
pubmed: 20658976
doi: 10.1042/BST0380914
Saar, K. L. et al. Rapid two-dimensional characterisation of proteins in solution. Microsyst. Nanoeng. 5, 1–10 (2019).
doi: 10.1038/s41378-019-0072-3
Emmenegger, M. et al. LAG3 is not expressed in human and murine neurons and does not modulate α-synucleinopathies. EMBO Mol. Med. 13, e14745 (2021).
pubmed: 34309222
pmcid: 8422075
doi: 10.15252/emmm.202114745
Sahtoe, D. D. et al. Design of amyloidogenic peptide traps. bioRxiv 2023.01.13.523785 https://doi.org/10.1101/2023.01.13.523785 (2023).
Hoyer, W. et al. Dependence of α-synuclein aggregate morphology on solution conditions. J. Mol. Biol. 322, 383–393 (2002).
pubmed: 12217698
doi: 10.1016/S0022-2836(02)00775-1
Herling, T. W. et al. Non-specificity fingerprints for clinical stage antibodies in solution. bioRxiv 2023.02.13.528263 https://doi.org/10.1101/2023.02.13.528263 (2023).
Duffy, D. C., McDonald, J. C., Schueller, O. J. A. & Whitesides, G. M. Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70, 4974–4984 (1998).
pubmed: 21644679
doi: 10.1021/ac980656z
Challa, P. K., Kartanas, T., Charmet, J. & Knowles, T. P. J. Microfluidic devices fabricated using fast wafer-scale LED-lithography patterning. Biomicrofluidics 11, 014113 (2017).
pubmed: 28289484
pmcid: 5315664
doi: 10.1063/1.4976690
Tan, S. H., Nguyen, N.-T., Chua, Y. C. & Kang, T. G. Oxygen plasma treatment for reducing hydrophobicity of a sealed polydimethylsiloxane microchannel. Biomicrofluidics 4, 032204 (2010).
pubmed: 21045926
pmcid: 2967237
doi: 10.1063/1.3466882