Abstract
Huntington’s disease is a neurodegenerative disorder caused by a CAG expansion in the first exon of the HTT gene, resulting in an extended polyglutamine (poly-Q) tract in huntingtin (httex1). The structural changes occurring to the poly-Q when increasing its length remain poorly understood due to its intrinsic flexibility and the strong compositional bias. The systematic application of site-specific isotopic labeling has enabled residue-specific NMR investigations of the poly-Q tract of pathogenic httex1 variants with 46 and 66 consecutive glutamines. Integrative data analysis reveals that the poly-Q tract adopts long α-helical conformations propagated and stabilized by glutamine side chain to backbone hydrogen bonds. We show that α-helical stability is a stronger signature in defining aggregation kinetics and the structure of the resulting fibrils than the number of glutamines. Our observations provide a structural perspective of the pathogenicity of expanded httex1 and pave the way to a deeper understanding of poly-Q-related diseases.
This is a preview of subscription content, access via your institution
Access options
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
Data availability
The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request. The accession codes for the SAXS data (SASDB) are SASDQR8 (H16) and SASDQS8 (H46) and the ensembles have been deposited in the Protein Ensemble Database under the accession codes PED00223 (H16) and PED00224 (H46). The 3D structure of sfGFP was downloaded from the PDB (PDB 3LVA). Source data are provided with this paper.
Code availability
The in-house script to analyze fluorescence images and disordered chain building program will be made available from the corresponding author on reasonable request.
References
Orr, H. T. Beyond the Qs in the polyglutamine diseases. Genes Dev. 15, 925–932 (2001).
Walker, F. O. Huntington’s disease. Lancet 369, 218–228 (2007).
Saudou, F. & Humbert, S. The biology of huntingtin. Neuron 89, 910–926 (2016).
Kremer, B. et al. A worldwide study of the Huntington’s disease mutation: the sensitivity and specificity of measuring CAG repeats. N. Engl. J. Med. 330, 1401–1406 (1994).
Benn, C. L. et al. Contribution of nuclear and extranuclear polyQ to neurological phenotypes in mouse models of Huntington’s disease. Hum. Mol. Genet. 14, 3065–3078 (2005).
Graham, R. K. et al. Cleavage at the caspase-6 site is required for neuronal dysfunction and degeneration due to mutant huntingtin. Cell 125, 1179–1191 (2006).
Zuccato, C., Valenza, M. & Cattaneo, E. Molecular mechanisms and potential therapeutical targets in Huntington’s disease. Physiol. Rev. 90, 905–981 (2010).
Mangiarini, L. et al. Exon 1 of the HD gene with an expanded CAG repeat is sufficient to cause a progressive neurological phenotype in transgenic mice. Cell 87, 493–506 (1996).
Feng, X., Luo, S. & Lu, B. Conformation polymorphism of polyglutamine proteins. Trends Biochem. Sci. 43, 424–435 (2018).
Caron, N. S., Desmond, C. R., Xia, J. & Truant, R. Polyglutamine domain flexibility mediates the proximity between flanking sequences in huntingtin. Proc. Natl Acad. Sci. USA 110, 14610–14615 (2013).
Nucifora, L. G. et al. Identification of novel potentially toxic oligomers formed in vitro from mammalian-derived expanded huntingtin exon-1 protein. J. Biol. Chem. 287, 16017–16028 (2012).
Li, P. et al. The structure of a polyQ–anti-polyQ complex reveals binding according to a linear lattice model. Nat. Struct. Mol. Biol. 14, 381–387 (2007).
Owens, G. E., New, D. M., West, A. P. & Bjorkman, P. J. Anti-polyQ antibodies recognize a short polyQ stretch in both normal and mutant huntingtin exon 1. J. Mol. Biol. 427, 2507–2519 (2015).
Warner, J. B. et al. Monomeric huntingtin exon 1 has similar overall structural features for wild-type and pathological polyglutamine lengths. J. Am. Chem. Soc. 139, 14456–14469 (2017).
Newcombe, E. A. et al. Tadpole-like conformations of huntingtin exon 1 are characterized by conformational heterogeneity that persists regardless of polyglutamine length. J. Mol. Biol. 430, 1442–1458 (2018).
Bravo-Arredondo, J. M. et al. The folding equilibrium of huntingtin exon 1 monomer depends on its polyglutamine tract. J. Biol. Chem. 293, 19613–19623 (2018).
Mier, P. et al. Disentangling the complexity of low complexity proteins. Brief. Bioinform. 21, 458–472 (2020).
Thakur, A. K. et al. Polyglutamine disruption of the huntingtin exon 1 N terminus triggers a complex aggregation mechanism. Nat. Struct. Mol. Biol. 16, 380–389 (2009).
Baias, M. et al. Structure and dynamics of the huntingtin exon-1 N-terminus: a solution NMR perspective. J. Am. Chem. Soc. 139, 1168–1176 (2017).
Ceccon, A. et al. Interaction of huntingtin exon-1 peptides with lipid-based micellar nanoparticles probed by solution NMR and Q-band pulsed EPR. J. Am. Chem. Soc. 140, 6199–6202 (2018).
Urbanek, A. et al. Site-specific isotopic labeling (SSIL): access to high-resolution structural and dynamic information in low-complexity proteins. ChemBioChem 21, 769–775 (2020).
Urbanek, A. et al. A general strategy to access structural information at atomic resolution in polyglutamine homorepeats. Angew. Chem. Int. Ed. Engl. 57, 3598–3601 (2018).
Urbanek, A. et al. Flanking regions determine the structure of the poly-glutamine in huntingtin through mechanisms common among glutamine-rich human proteins. Structure 28, 733–746.e5 (2020).
Shen, K. et al. Control of the structural landscape and neuronal proteotoxicity of mutant huntingtin by domains flanking the polyQ tract. eLife 5, e18065 (2016).
Bhattacharyya, A. et al. Oligoproline effects on polyglutamine conformation and aggregation. J. Mol. Biol. 355, 524–535 (2006).
Morató, A. et al. Robust cell-free expression of sub-pathological and pathological huntingtin exon-1 for NMR studies. General approaches for the isotopic labeling of low-complexity proteins. Biomolecules 10, 1458 (2020).
Nielsen, J. T. & Mulder, F. A. A. POTENCI: prediction of temperature, neighbor and pH-corrected chemical shifts for intrinsically disordered proteins. J. Biomol. NMR 70, 141–165 (2018).
Milles, S., Salvi, N., Blackledge, M. & Jensen, M. R. Characterization of intrinsically disordered proteins and their dynamic complexes: from in vitro to cell-like environments. Prog. Nucl. Magn. Reson. Spectrosc. 109, 79–100 (2018).
Estaña, A. et al. Realistic ensemble models of intrinsically disordered proteins using a structure-encoding coil database. Structure 27, 381–391 (2019).
Tria, G., Mertens, H. D. T., Kachala, M. & Svergun, D. I. Advanced ensemble modelling of flexible macromolecules using X-ray solution scattering. IUCrJ 2, 207–217 (2015).
Bernadó, P., Mylonas, E., Petoukhov, M. V., Blackledge, M. & Svergun, D. I. Structural characterization of flexible proteins using small-angle X-ray scattering. J. Am. Chem. Soc. 129, 5656–5664 (2007).
Iglesias, J., Sanchez-Martínez, M. & Crehuet, R. SS-map: visualizing cooperative secondary structure elements in protein ensembles. Intrinsically Disord. Proteins 1, e25323 (2013).
Escobedo, A. et al. Side chain to main chain hydrogen bonds stabilize a polyglutamine helix in a transcription factor. Nat. Commun. 10, 2034 (2019).
Qu, W. et al. Synthesis of optically pure 4-fluoro-glutamines as potential metabolic imaging agents for tumors. J. Am. Chem. Soc. 133, 1122–1133 (2011).
Gimenez, D., Phelan, A., Murphy, C. D. & Cobb, S. L. 19F NMR as a tool in chemical biology. Beilstein J. Org. Chem. 17, 293–318 (2021).
Kitevski-LeBlanc, J. L. & Prosser, R. S. Current applications of 19F NMR to studies of protein structure and dynamics. Prog. Nucl. Magn. Reson. Spectrosc. 62, 1–33 (2012).
Miao, Y., Feher, V. A. & McCammon, J. A. Gaussian accelerated molecular dynamics: unconstrained enhanced sampling and free energy calculation. J. Chem. Theory Comput. 11, 3584–3595 (2015).
Tang, W. S., Fawzi, N. L. & Mittal, J. Refining all-atom protein force fields for polar-rich, prion-like, low-complexity intrinsically disordered proteins. J. Phys. Chem. B 124, 9505–9512 (2020).
Monsellier, E., Redeker, V., Ruiz-Arlandis, G., Bousset, L. & Melki, R. Molecular interaction between the chaperone Hsc70 and the N-terminal flank of huntingtin exon 1 modulates aggregation. J. Biol. Chem. 290, 2560–2576 (2015).
Galaz-Montoya, J. G., Shahmoradian, S. H., Shen, K., Frydman, J. & Chiu, W. Cryo-electron tomography provides topological insights into mutant huntingtin exon 1 and PolyQ aggregates. Commun. Biol. 4, 849 (2021).
Ruggeri, F. S. et al. Nanoscale studies link amyloid maturity with polyglutamine diseases onset. Sci. Rep. 6, 31155 (2016).
Ormsby, A. R. et al. Nascent mutant huntingtin exon 1 chains do not stall on ribosomes during translation but aggregates do recruit machinery involved in ribosome quality control and RNA. PLoS ONE 15, e0233583 (2020).
Vieweg, S. et al. The Nt17 domain and its helical conformation regulate the aggregation, cellular properties and neurotoxicity of mutant huntingtin exon 1. J. Mol. Biol. 433, 167222 (2021).
Riguet, N. et al. Nuclear and cytoplasmic huntingtin inclusions exhibit distinct biochemical composition, interactome and ultrastructural properties. Nat. Commun. 12, 6579 (2021).
Bäuerlein, F. J. B. et al. In situ architecture and cellular interactions of PolyQ inclusions. Cell 171, 179–187.e10 (2017).
Moldovean, S. N. & Chiş, V. Molecular dynamics simulations applied to structural and dynamical transitions of the huntingtin protein: a review. ACS Chem. Neurosci. 11, 105–120 (2020).
Kang, H. et al. Emerging β-sheet rich conformations in supercompact huntingtin exon-1 mutant structures. J. Am. Chem. Soc. 139, 8820–8827 (2017).
Kotler, S. A. et al. Probing initial transient oligomerization events facilitating huntingtin fibril nucleation at atomic resolution by relaxation-based NMR. Proc. Natl Acad. Sci. USA 116, 3562–3571 (2019).
Wetzel, R. Exploding the repeat length paradigm while exploring amyloid toxicity in Huntington’s disease. Acc. Chem. Res. 53, 2347–2357 (2020).
Drombosky, K. W. et al. Mutational analysis implicates the amyloid fibril as the toxic entity in Huntington’s disease. Neurobiol. Dis. 120, 126–138 (2018).
Sathasivam, K. et al. Identical oligomeric and fibrillar structures captured from the brains of R6/2 and knock-in mouse models of Huntington’s disease. Hum. Mol. Genet. 19, 65–78 (2010).
Gruber, A. et al. Molecular and structural architecture of polyQ aggregates in yeast. Proc. Natl Acad. Sci. USA 115, E3446–E3453 (2018).
Mario Isas, J. et al. Huntingtin fibrils with different toxicity, structure, and seeding potential can be interconverted. Nat. Commun. 12, 4272 (2021).
Ceccon, A., Tugarinov, V., Ghirlando, R. & Clore, G. M. Abrogation of prenucleation, transient oligomerization of the huntingtin exon 1 protein by human profilin I. Proc. Natl Acad. Sci. USA 117, 5844–5852 (2020).
Fiumara, F., Fioriti, L., Kandel, E. R. & Hendrickson, W. A. Essential role of coiled coils for aggregation and activity of Q/N-rich prions and polyQ proteins. Cell 143, 1121–1135 (2010).
Jayaraman, M. et al. Kinetically competing huntingtin aggregation pathways control amyloid polymorphism and properties. Biochemistry 51, 2706–2716 (2012).
Nekooki-Machida, Y. et al. Distinct conformations of in vitro and in vivo amyloids of huntingtin-exon1 show different cytotoxicity. Proc. Natl Acad. Sci. USA 106, 9679–9684 (2009).
Rockabrand, E. et al. The first 17 amino acids of huntingtin modulate its sub-cellular localization, aggregation and effects on calcium homeostasis. Hum. Mol. Genet. 16, 61–77 (2007).
Atwal, R. S. & Truant, R. A stress sensitive ER membrane-association domain in huntingtin protein defines a potential role for huntingtin in the regulation of autophagy. Autophagy 4, 91–93 (2008).
Atwal, R. S. et al. Huntingtin membrane association signal that can modulate huntingtin aggregation, nuclear entry and toxicity. Hum. Mol. Genet. 16, 2600–2615 (2007).
Michalek, M., Salnikov, E. S., Werten, S. & Bechinger, B. Membrane interactions of the amphipathic amino terminus of huntingtin. Biochemistry 52, 847–858 (2013).
Marquette, A., Aisenbrey, C. & Bechinger, B. Membrane interactions accelerate the self-aggregation of huntingtin exon 1 fragments in a polyglutamine length-dependent manner. Int. J. Mol. Sci. 22, 6725 (2021).
Michalek, M., Salnikov, E. S. & Bechinger, B. Structure and topology of the huntingtin 1–17 membrane anchor by a combined solution and solid-state NMR approach. Biophys. J. 105, 699–710 (2013).
Loscha, K. V. et al. Multiple-site labeling of proteins with unnatural amino acids. Angew. Chem. Int. Ed. Engl. 51, 2243–2246 (2012).
Apponyi, M. A., Ozawa, K., Dixon, N. E. & Otting, G. in Structural Proteomics: High-Throughput Methods (eds Kobe, B. et al.) 257–268 (Humana Press, 2008).
Vranken, W. F. et al. The CCPN data model for NMR spectroscopy: development of a software pipeline. Proteins 59, 687–696 (2005).
Markley, J. L. et al. Recommendations for the presentation of NMR structures of proteins and nucleic acids. J. Mol. Biol. 280, 933–952 (1998).
Andreeva, A., Howorth, D., Chothia, C., Kulesha, E. & Murzin, A. G. SCOP2 prototype: a new approach to protein structure mining. Nucleic Acids Res. 42, D310–D314 (2014).
Andreeva, A., Kulesha, E., Gough, J. & Murzin, A. G. The SCOP database in 2020: expanded classification of representative family and superfamily domains of known protein structures. Nucleic Acids Res. 48, D376–D382 (2020).
Krivov, G. G., Shapovalov, M. V. & Dunbrack Jr., R. L. Improved prediction of protein side-chain conformations with SCWRL4. Proteins 77, 778–795 (2009).
Shen, Y. & Bax, A. SPARTA+: a modest improvement in empirical NMR chemical shift prediction by means of an artificial neural network. J. Biomol. NMR 48, 13–22 (2010).
Thureau, A., Roblin, P. & Pérez, J. BioSAXS on the SWING beamline at Synchrotron SOLEIL. J. Appl. Crystallogr. 54, 1698–1710 (2021).
Blanchet, C. E. et al. Versatile sample environmentsand automation for biological solution X-ray scattering experiments at the P12 Beamline (PETRA III, DESY). J. Appl. Crystallogr. 48, 431–443 (2015).
Girardot, R., Viguier, G., Pérez, J. & Ounsy, M. FOXTROT: a Java-based application to reduce and analyse SAXS and WAXS piles of 2D data at Synchrotron SOLEIL. In Proc. canSAS-VIII (2015).
Panjkovich, A. & Svergun, D. I. CHROMIXS: automatic and interactive analysis of chromatography-coupled small-angle X-ray scattering data. Bioinformatics 34, 1944–1946 (2018).
Franke, D. et al. ATSAS 2.8: a comprehensive data analysis suite for small-angle scattering from macromolecular solutions. J. Appl. Crystallogr. 50, 1212–1225 (2017).
Svergun, D. I. Determination of the regularization parameter in indirect-transform methods using perceptual criteria. J. Appl. Crystallogr. 25, 495–503 (1992).
Hajizadeh, N. R., Franke, D., Jeffries, C. M. & Svergun, D. I. Consensus Bayesian assessment of protein molecular mass from solution X-Ray scattering data. Sci. Rep. 8, 7204 (2018).
Humphrey, W., Dalke, A. & Schulten, K. VMD: visual molecular dynamics. J. Mol. Graph. 14, 33–38 (1996).
Abraham, M. J. et al. GROMACS: high performance molecular simulations through multi-level parallelism from laptops to supercomputers. SoftwareX 1–2, 19–25 (2015).
Best, R. B., Zheng, W. & Mittal, J. Balanced protein–water interactions improve properties of disordered proteins and non-specific protein association. J. Chem. Theory Comput. 10, 5113–5124 (2014).
Shirts, M. R. et al. Lessons learned from comparing molecular dynamics engines on the SAMPL5 dataset. J. Comput. Aided Mol. Des. 31, 147–161 (2017).
Case, D. A. et al. AMBER 2016 (Univ. California, San Francisco, 2016).
Chow, K.-H. & Ferguson, D. M. Isothermal-isobaric molecular dynamics simulations with Monte Carlo volume sampling. Comput. Phys. Commun. 91, 283–289 (1995).
Neese, F., Wennmohs, F., Becker, U. & Riplinger, C. The ORCA quantum chemistry program package. J. Chem. Phys. 152, 224108 (2020).
Neese, F. Software update: the ORCA program system—version 5.0. Wiley Interdiscip. Rev. Comput. Mol. Sci. 12, e1606 (2022).
Field, M. J. The pDynamo program for molecular simulations using hybrid quantum chemical and molecular mechanical potentials. J. Chem. Theory Comput. 4, 1151–1161 (2008).
Tao, J., Perdew, J. P., Staroverov, V. N. & Scuseria, G. E. Climbing the density functional ladder: nonempirical meta-generalized gradient approximation designed for molecules and solids. Phys. Rev. Lett. 91, 146401 (2003).
Stoychev, G. L., Auer, A. A., Izsák, R. & Neese, F. Self-consistent field calculation of nuclear magnetic resonance chemical shielding constants using gauge-including atomic orbitals and approximate two-electron integrals. J. Chem. Theory Comput. 14, 619–637 (2018).
Schattenberg, C. J. & Kaupp, M. Extended benchmark set of main-group nuclear shielding constants and NMR chemical shifts and its use to evaluate modern DFT methods. J. Chem. Theory Comput. 17, 7602–7621 (2021).
Grimme, S., Ehrlich, S. & Goerigk, L. Effect of the damping function in dispersion corrected density functional theory. J. Comput. Chem. 32, 1456–1465 (2011).
Grimme, S., Antony, J., Ehrlich, S. & Krieg, H. A consistent and accurate ab initio parametrization of density functional dispersion correction (DFT-D) for the 94 elements H-Pu. J. Chem. Phys. 132, 154104 (2010).
Jensen, F. Segmented contracted basis sets optimized for nuclear magnetic shielding. J. Chem. Theory Comput. 11, 132–138 (2015).
Cammi, R., Mennucci, B. & Tomasi, J. Fast evaluation of geometries and properties of excited molecules in solution: a Tamm-Dancoff model with application to 4-dimethylaminobenzonitrile. J. Phys. Chem. A 104, 5631–5637 (2000).
Fedorov, S. V. & Krivdin, L. B. Computational protocols for the 19F NMR chemical shifts. Part 1: methodological aspects. J. Fluor. Chem. 238, 109625 (2020).
Dahmane, S. et al. Nanoscale organization of tetraspanins during HIV-1 budding by correlative dSTORM/AFM. Nanoscale 11, 6036–6044 (2019).
Proksch, R., Schäffer, T. E., Cleveland, J. P., Callahan, R. C. & Viani, M. B. Finite optical spot size and position corrections in thermal spring constant calibration. Nanotechnology 15, 1344–1350 (2004).
Acknowledgements
We thank G. Otting (Australian National University, Canberra, Australia) for providing the BL21 (DE3) Star::RF1-CBD3 strain. This work was supported by the European Research Council under the European Union’s H2020 Framework Programme (2014–2020)/ERC grant agreement no. 648030 and Labex EpiGenMed, an Investissements d’avenir program (grant no. ANR-10-LABX-12-01) awarded to P.B., grant no. ANR-17-CE11-0022-01 awarded to N.S. and grant no. ANR-19-PI3A-0004 awarded to J.C. The Centre for Structural Biology (CBS) is a member of France-BioImaging (FBI) and the French Infrastructure for Integrated Structural Biology, two national infrastructures supported by the French National Research Agency (grant nos. ANR-10-INBS-04-01 and ANR-10-INBS-05, respectively). A.U. is supported by a grant from the Fondation pour la Recherche Médicale (grant no. SPF20150934061). D.S. acknowledges a grant from the Métropole Européenne de Lille (PUSHUP). G. Levy (Université de Lille) is thanked for help with sample preparation and the 19F-NMR experiments. This work benefited from the high-performance computing resources of CSUC and the CALMIP supercomputing center under the allocations 2016-P16032 and 2021-P21043. The 600 MHz spectrometer for 19F-NMR measurements is funded by the Nord Region Council, CNRS, Institut Pasteur de Lille, the European Community (European Regional Development Fund, ERDF), the French Ministry of Research and the Université de Lille and by the CTRL CPER cofunded by the European Union with the ERDF, by the Hauts-de-France Regional Council (contract no. 17003781), Métropole Européenne de Lille (contract no. 2016_ESR_05) and the French State (contract no .2017-R3-CTRL-Phase1). We thank the SWING beamline at the SOLEIL synchrotron, Saint-Aubin, France (proposal 20181386), and P12 beamline at PETRAIII, Hamburg, Germany, for beamtime allocation to the project and assistance during data collection.
Author information
Authors and Affiliations
Contributions
P.B. conceived the project. C.A.E.-R., A.U., A.S., C.D., R.C., A.B., D.S., N.S. and P.B. designed experiments. C.A.E.-R., A.U., A.S., M.P., A.M., A.E., A.F., C.D., X.L.L., Z.-D.S., L.C., A.T., F.A., R.C. and D.S. performed experiments. R.E.S., P.-E.M., A.B., J.C., N.S. and P.B. supervised experiments. C.A.E.-R., A.U., A.S. and P.B. wrote the paper with the help of all the coauthors.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests.
Peer review
Peer review information
Nature Structural & Molecular Biology thanks Burkhard Bechinger and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
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 Comparison of H46 and H16 by NMR.
(a) Overlay of the 15N-HSQC spectra of fully labeled H16 (red) with selectively labeled samples of H46 (blue). (b) Zoom of the 15N-HSQC and 13C-HSQC overlay of H16 and H46 SSIL spectra showing the poly-Q NH, NH and Cα regions of Q18, Q20 and Q21.
Extended Data Fig. 2 Comparison of H66 with H16 and H46 by NMR.
(Left) Overlay of the 15N-HSQC spectra of fully labeled H66 (green) with fully labeled H46 (blue). Black arrows indicate peaks corresponding to glutamates, which were not labeled in the H66 sample. (Right) Comparison of the SCS profiles of H16 (red), H46 (blue) and the values measured for Q56 and Q76 in H66 (green).
Extended Data Fig. 3 SAXS analyses of H46 and H16.
The plots of average intensity vs. frame number obtained from SEC–SAXS for H16 (a) and H46 (b) The insets show all−vs-all χ2 comparison for the frames selected for further processing. The small values of χ2 show that the selected frames were very similar to each other. (c) Pairwise distance distribution functions, P(r), obtained for H16 (red) and H46 (blue) by indirect Fourier transformation of the SAXS data. (d) and (e) show the Rg distribution of the pool (gray, filled) and the selected sub-ensemble by EOM for H16 (red) and H46 (blue), respectively.
Extended Data Fig. 5 Labeling of httex1 with fluorinated glutamines.
(a) Enzymatic loading of suppressor tRNACUA with canonical (Gln) or 2 S,4R-fluoroglutamine (4F-Gln). Upper and lower bands correspond to loaded and unloaded suppressor tRNACUA, respectively. A negative control of empty tRNACUA is shown in the third lane. (b) Averaged fluorescence endpoint derived from n = 2 independent cell-free suppression reactions of H16 with a stop codon at position Q18 or Q21 when titrating with increasing concentrations of both Gln-tRNACUA and 4F-Gln-tRNACUA. “+” indicates a positive control, a cell-free reaction of H16 without any amber stop codon. Data are presented as mean values + /- standard deviation.
Extended Data Fig. 6 Helicity propagation in H46 MD trajectories.
(a) Plot showing the number of times (in log scale) an α-helix expands 1 to 5 residues towards the C-terminus (+1 to +5) or the N-terminus (-1 to -5) from each of the residues of N17 and poly-Q. For all the positions (1 to 5), the number of events is greater in direction of the C-terminus than the N-terminus, indicating a higher propensity of helix propagation from N- to C-terminus. The comparison of the number of events for expansion of a helix by 1, 2 and 3 residues towards the N- or C-terminus is presented in (b) for further clarity. These plots show that in httex1 helical segments are more prone to expand towards the C-terminus than towards the N-terminus. Additionally, the preference for directional growing increases with the extent of propagation, that is, the relative difference in the number of events where a helix is propagated three residues towards the C-terminus and the N-terminus is larger than propagation the of two residues, which is in turn larger than the propagation of a single residue.
Extended Data Fig. 7 Quantum chemistry calculations of the 19F chemical shifts.
(A) Box plots of the 19F CS when the F-Gln was incorporated in Q20 (blue) or in Q21 (orange) for n = 50 distinct conformations belonging to the following families: (left, bifurcated) an α-helical structure with either a S16-Q20 or a F17-Q21 bifurcated hydrogen bond, (middle, non-bifurcated) having a canonical α-helical structure in the S16-Q21 segment without a bifurcated hydrogen bond, and (right, random coil) not displaying any specific secondary structure. A statistically significant difference was observed between the 19F CS distributions for Q20 and Q21 obtained from conformations with a bifurcated hydrogen bond between S16-Q20 and F17-Q21, respectively (p = 0.003). Q20 and Q21 19F chemical shifts were also statistically distinct when both residues were involved in an α-helix (p = 0.0023). No statistical difference was observed in the random coil scenario (p = 1.0). The boxplots use the default settings in the seaborn statistical plotting library. The centre of each box represents the median. The box limits represent the interquartile range, that is Q3-Q1. “Whiskers” extend to points that lie within 1.5 interquartile ranges of the lower and upper quartile. Observations that fall outside this range are displayed independently. The test used was the two-sided T-test for 2 independent samples with the same population variance with the Bonferroni correction, as implemented in the statannotation python library (https://github.com/trevismd/statannotations). (B) Cartoon displaying the distance between the fluorine atom when placed in Q21 and the center of the phenylalanine ring. (C) Influence of the distance on the computed 19F CSs of F-Q21 when adopting an α-helical conformation in the presence (blue) or absence (red) of a bifurcate hydrogen bond. Horizontal lines display the largest 19F CS values for the different scenarios. (D) Distance distribution between the fluorine atom when placed in Q21 and the center of the phenylalanine ring for the bifurcated and non-bifurcated hydrogen bonds computed from conformations extracted from the GaMD trajectory. In both cases, a large population of conformations placed the fluorine atom in very close proximity (<4 Å) of F17 aromatic ring.
Extended Data Fig. 8 NMR investigation of the N17 mutants exhibiting a different poly-Q helical propensity while preserving its length.
Uniformly labeled and SSIL samples of LKGG- and LLLF-H46 mutants were produced and analyzed by NMR. On one hand, LKGG-H46 glutamine 15N-HSQC signals (blue) collapsed in a broad, high-intensity, downfield-shifted peak, proving a substantial loss of helicity in comparison with the wild-type H46 (in gray) (a). Interestingly, this broad peak did not overlap with the positions corresponding to fully unstructured glutamines, which were shifted further downfield. This indicates that poly-Q, even when disconnected from the flanking region, contains a small intrinsic propensity for helical conformations, in agreement with our MD simulations. On the other hand, the 15N-HSQC spectrum of fully labeled LLLF-H46 (green) displayed a more dispersed density of glutamine peaks and an additional upfield density compared with the wild-type (in gray), pointing to a helicity increase of the poly-Q tract (b). The detailed analysis of NH, NεH2 and Cα signals of Q18, Q20 and Q21 SSIL samples from both mutants confirmed the decrease in structuration of LKGG-H46 and the increase in helicity in LLLF-H46, in comparison with the wild-type form (in gray) (a,b and c). Importantly, Q21 in LLLF-H46 displayed signatures of two conformations as also observed for the wild-type. This residue exhibited two NHε correlation peaks, suggesting the formation of a stronger bifurcated hydrogen bond than in the wild-type. Unfortunately, a single Cα peak, corresponding to the less helical conformation, was observed for Q21 in this mutant, suggesting an unfavorable exchange regime for the NMR detection.
Extended Data Fig. 9 AFM investigation of huntingtin fibrils.
(a) Fluorescence microscopy (upper panels) and AFM (middle and lower panels) images of 2-day-old fibrils of H46, LLLF-H46 and LKGG-H46. Each fluorescence image corresponds to the average of 150 pictures. (b) AFM images of 5-day-old fibrils of the three H46 variants. White squares indicate the zoom region displayed in panels below. At least 6 fluorescence and AFM images for each H46 variant and time point were extracted from two large fields (8.13μm x 8.13μm). Images displayed in (a) and (b) are representative of the ensemble of them.
Supplementary information
Supplementary Information
Supplementary Figs. 1–6, Notes 1 and 2, Discussion and Tables 1–4.
Source data
Source Data Extended Data Fig. 5
Unprocessed urea PAGE gel to monitor the loading of F-Gln onto tRNA.
Rights and permissions
Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.
About this article
Cite this article
Elena-Real, C.A., Sagar, A., Urbanek, A. et al. The structure of pathogenic huntingtin exon 1 defines the bases of its aggregation propensity. Nat Struct Mol Biol 30, 309–320 (2023). https://doi.org/10.1038/s41594-023-00920-0
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41594-023-00920-0
This article is cited by
-
Alternative low-populated conformations prompt phase transitions in polyalanine repeat expansions
Nature Communications (2024)
-
The molecular basis for cellular function of intrinsically disordered protein regions
Nature Reviews Molecular Cell Biology (2024)