Abstract
The small heat shock protein αA-crystallin is a molecular chaperone important for the optical properties of the vertebrate eye lens. It forms heterogeneous oligomeric ensembles. We determined the structures of human αA-crystallin oligomers by combining cryo-electron microscopy, cross-linking/mass spectrometry, NMR spectroscopy and molecular modeling. The different oligomers can be interconverted by the addition or subtraction of tetramers, leading to mainly 12-, 16- and 20-meric assemblies in which interactions between N-terminal regions are important. Cross-dimer domain-swapping of the C-terminal region is a determinant of αA-crystallin heterogeneity. Human αA-crystallin contains two cysteines, which can form an intramolecular disulfide in vivo. Oxidation in vitro requires conformational changes and oligomer dissociation. The oxidized oligomers, which are larger than reduced αA-crystallin and destabilized against unfolding, are active chaperones and can transfer the disulfide to destabilized substrate proteins. The insight into the structure and function of αA-crystallin provides a basis for understanding its role in the eye lens.
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
Rent or buy this article
Prices vary by article type
from$1.95
to$39.95
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
Data availability
The cryo-EM density maps of αA-crystallin oligomers have been deposited in the EMBD under accession codes EMD-4895 (12-mer), EMD-4894 (16-mer) and EMD-4896 (20-mer). The coordinates for the 16-mer model were deposited in the wwPDB under accession number PDB 6T1R. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with dataset identifier PXD013587. The1H, 15N and 13C chemical shifts of reduced αA-crystallin are available at the BioMagResBank (BMRB) with accession number BMRB-27109. All other data are available from the corresponding authors upon reasonable request.
References
Bloemendal, H. et al. Ageing and vision: structure, stability and function of lens crystallins. Prog. Biophys. Mol. Biol. 86, 407–485 (2004).
Clark, A. R., Lubsen, N. H. & Slingsby, C. sHSP in the eye lens: crystallin mutations, cataract and proteostasis. Int. J. Biochem. Cell Biol. 44, 1687–1697 (2012).
Horwitz, J. α-Crystallin can function as a molecular chaperone. Proc. Natl Acad. Sci. USA 89, 10449–10453 (1992).
Horwitz J. Alpha-crystallin. Exp. Eye Res. 76, 145–153 (2003).
Graw, J. Genetics of crystallins: cataract and beyond. Exp. Eye Res. 88, 173–189 (2009).
Oguni, M. et al. Ontogeny of alpha-crystallin subunits in the lens of human and rat embryos. Cell Tissue Res. 276, 151–154 (1994).
Iwaki, T., Kume-Iwaki, A., Liem, R. K. H. & Goldman, J. E. αB-crystallin is expressed in non-lenticular tissues and accumulates in Alexander’s disease brain. Cell 57, 71–78 (1989).
Srinivasan, A. N., Nagineni, C. N. & Bhat, S. P. αA-crystallin is expressed in non-ocular tissues. J. Biol. Chem. 267, 23337–23341 (1992).
Gangalum, R. K., Horwitz, J., Kohan, S. A. & Bhat, S. P. αA-crystallin and αB-crystallin reside in separate subcellular compartments in the developing ocular lens. J. Biol. Chem. 287, 42407–42416 (2012).
Datta, S. A. & Rao, C. M. Differential temperature-dependent chaperone-like activity of αA- and αB-crystallin homoaggregates. J. Biol. Chem. 274, 34773–34778 (1999).
Reddy, G. B., Das, K. P., Petrash, J. M. & Surewicz, W. K. Temperature-dependent chaperone activity and structural properties of human αA- and αB-crystallins. J. Biol. Chem. 275, 4565–4570 (2000).
Kumar, L. V., Ramakrishna, T. & Rao, C. M. Structural and functional consequences of the mutation of a conserved arginine residue in αA- and αB-crystallins. J. Biol. Chem. 274, 24137–24141 (1999).
Caspers, G. J., Leunissen, J. A. & de Jong, W. W. The expanding small heat-shock protein family, and structure predictions of the conserved ‘α-crystallin domain’. J. Mol. Evol. 40, 238–248 (1995).
de Jong, W. W., Caspers, G. J. & Leunissen, J. A. Genealogy of the α-crystallin—small heat-shock protein superfamily. Int. J. Biol. Macromol. 22, 151–162 (1998).
Laganowsky, A. et al. Crystal structures of truncated alphaA and alphaB crystallins reveal structural mechanisms of polydispersity important for eye lens function. Protein Sci. 19, 1031–1043 (2010).
Laganowsky, A. & Eisenberg, D. Non-3D domain swapped crystal structure of truncated zebrafish alphaA crystallin. Protein Sci. 19, 1978–1984 (2010).
Bova, M. P., Ding, L. L., Horwitz, J. & Fung, B. K. Subunit exchange of αA-crystallin. J. Biol. Chem. 272, 29511–29517 (1997).
Aquilina, J. A. et al. Subunit exchange of polydisperse proteins: mass spectrometry reveals consequences of αA-crystallin truncation. J. Biol. Chem. 280, 14485–1449 (2005).
Peschek, J. et al. The eye lens chaperone α-crystallin forms defined globular assemblies. Proc. Natl Acad. Sci. USA 106, 13272–13277 (2009).
Merck, K. B., de Haard-Hoekman, W. A., Essink, B. B. O., Bloemendal, H. & de Jong, W. W. Expression and aggregation of recombinant αA-crystallin and its two domains. Biochim. Biophys. Acta 1130, 267–276 (1992).
Bova, M. P., McHaourab, H. S., Han, Y. & Fung, B. K. K. Subunit exchange of small heat shock proteins. Analysis of oligomer formation of αA-crystallin and Hsp27 by fluorescence resonance energy transfer and site-directed truncations. J. Biol. Chem. 275, 1035–1042 (2000).
Salerno, J. C., Eifert, C. L., Salerno, K. M. & Koretz, J. F. Structural diversity in the small heat shock protein superfamily: control of aggregation by the N-terminal region. Protein Eng. 16, 847–851 (2003).
Kundu, M., Sen, P. C. & Das, K. P. Structure, stability and chaperone function of αA-crystallin: role of N-terminal region. Biopolymers 86, 177–192 (2007).
Andley, U. P., Mathur, S., Griest, T. A. & Petrash, J. M. Cloning, expression and chaperone-like activity of human αA-crystallin. J. Biol. Chem. 271, 31973–31980 (1996).
Thampi, P. & Abraham, E. C. Influence of the C-terminal residues on oligomerization of αA-crystallin. Biochemistry 42, 11857–11863 (2003).
Rajan, S., Chandrashekar, R., Aziz, A. & Abraham, E. C. Role of arginine-163 and the 163REEK166 motif in the oligomerization of truncated α-crystallins. Biochemistry 45, 15684–15691 (2006).
Aziz, A., Santhoshkumar, P., Sharma, K. K. & Abraham, E. C. Cleavage of the C-terminal serine of human αA-crystallin produces αA1–172 with increased chaperone activity and oligomeric size. Biochemistry 46, 2510–2519 (2007).
Treweek, T. M., Rekas, A., Walker, M. J. & Carver, J. A. A quantitative NMR spectroscopic examination of the flexibility of the C-terminal extensions of the molecular chaperones, αA- and αB-crystallin. Exp. Eye Res. 91, 691–699 (2010).
Kim, K. K., Kim, R. & Kim, S. H. Crystal structure of a small heat-shock protein. Nature 394, 595–599 (1998).
van Montfort, R. L., Basha, E., Friedrich, K. L., Slingsby, C. & Vierling, E. Crystal structure and assembly of a eukaryotic small heat shock protein. Nat. Struct. Biol. 8, 1025–1030 (2001).
Runkle, S., Hill, J., Kantorow, M., Horwitz, J. & Posner, M. Sequence and spatial expression of zebrafish (Danio rerio) αA-crystallin. Mol. Vis. 8, 45–50 (2002).
Augusteyn, R. C., Hum, T. P., Putilin, T. P. & Thomson, J. A. The location of sulphydryl groups in α-crystallin. Biochim. Biophys. Acta 915, 132–139 (1987).
Srikanthan, D., Bateman, O. A., Purkiss, A. G. & Slingsby, C. Sulfur in human crystallins. Exp. Eye Res. 79, 823–831 (2004).
Miesbauer, L. R. et al. Post-translational modifications of water-soluble human lens crystallins from young adult. J. Biol. Chem. 269, 12494–12502 (1994).
Takemoto, L. J. Oxidation of cysteine residues from alpha-A crystallin during cataractogenesis of the human lens. Biochem. Biophys. Res. Commun. 223, 216–220 (1996).
Takemoto, L. J. Increase in the intramolecular disulfide bonding of alpha-A crystallin during aging of the human lens. Exp. Eye Res. 63, 585–590 (1996).
Hains, P. G. & Truscott, R. J. W. Proteomic analysis of the oxidation of cysteine residues in human age-related nuclear cataract lenses. Biochim. Biophys. Acta. 1784, 1959–1964 (2008).
Fan, X. et al. Evidence of highly conserved β-crystallin disulfidome that can be mimicked by in vitro oxidation in age-related human cataract and glutathione depleted mouse lens. Mol. Cell. Proteomics 14, 3211–3223 (2015).
Yang, Z., Chamorro, M., Smith, D. L. & Smith, J. B. Identification of the major components of the high molecular weight crystallins from old human lenses. Curr. Eye Res. 13, 415–421 (1994).
Lund, A. L., Smith, J. B. & Smith, D. L. Modifications of the water-insoluble human lens α-crystallins. Exp. Eye Res. 63, 661–672 (1996).
Hanson, S. R. A., Hasan, A., Smith, D. L. & Smith, J. B. The major in vivo modifications of the human water-insoluble lens crystallins are disulfide bonds, deamidation, methionine oxidation and backbone cleavage. Exp. Eye Res. 71, 195–207 (2000).
Cherian-Shaw, M., Smith, J. B., Jiang, X. Y. & Abraham, E. C. Intrapolypeptide disulfides in human αA-crystallin and their effect on chaperone-like function. Mol. Cell. Biochem. 199, 163–167 (1999).
Merkley, E. D. et al. Distance restraints from crosslinking mass spectrometry: mining a molecular dynamics simulation database to evaluate lysine–lysine distances. Protein Sci. 23, 747–759 (2014).
Dyson, H. J. & Wright, P. E. Unfolded proteins and protein folding studied by NMR. Chem. Rev. 104, 3607–3622 (2004).
Clore, G. M. & Iwahara, J. Theory, practice and applications of paramagnetic relaxation enhancement for the characterization of transient low-population states of biological macromolecules and their complexes. Chem. Rev. 109, 4108–4139 (2009).
Chakraborty, K. et al. Protein stabilization by introduction of cross-strand disulfides. Biochemistry 44, 14638–14646 (2005).
Wunderlich, M. & Glockshuber, R. Redox properties of protein disulfide isomerase (DsbA) from Escherichia coli. Protein Sci. 2, 717–726 (1993).
Zapun, A., Missiakas, D., Raina, S. & Creighton, T. E. Structural and functional characterization of DsbC, a protein involved in disulfide bond formation in Escherichia coli. Biochemistry 34, 5075–5089 (1995).
Hasan, A., Yu, J., Smith, D. L. & Smith, J. B. Thermal stability of human α-crystallins sensed by amide hydrogen exchange. Protein Sci. 13, 332–341 (2004).
Jehle, S. et al. N-terminal domain of αB-crystallin provides a conformational switch for multimerization and structural heterogeneity. Proc. Natl Acad. Sci. USA 108, 6409–6414 (2011).
Mainz, A. et al. The chaperone αB-crystallin uses different interfaces to capture an amorphous and an amyloid client. Nat. Struct. Mol. Biol. 22, 898–905 (2015).
Sluchanko, N. N. et al. Structural basis for the interaction of a human small heat shock protein with the 14–3–3 universal signaling regulator. Structure 25, 305–316 (2017).
Pasta, Y., Raman, B., Ramakrishna, T. & Rao, C. M. Role of the conserved SRLFDQFFG region of α-crystallin, a small heat shock protein. Effect on oligomeric size, subunit exchange and chaperone-like activity. J. Biol. Chem. 278, 51159–51166 (2003).
Baldwin, A. et al. Quaternary dynamics of αB-crystallin as a direct consequence of localised tertiary fluctuations in the C-terminus. J. Mol. Biol. 413, 310–320 (2011).
Alderson, T. R., Benesch, J. L. P. & Baldwin, A. J. Proline isomerization in the C-terminal region of HSP27. Cell Stress Chaperones 22, 639–651 (2017).
Jehle, S. et al. Solid-state NMR and SAXS studies provide a structural basis for the activation of αB-crystallin oligomers. Nat. Struct. Mol. Biol. 17, 1037–1042 (2010).
Braun, N. et al. Multiple molecular architectures of the eye lens chaperone αB-crystallin elucidated by a triple hybrid approach. Proc. Natl Acad. Sci. USA 108, 20491–20496 (2011).
Pasta, S. Y., Raman, B., Ramakrishna, T. & Rao, C. M. The IXI/V motif in the C-terminal extension of α-crystallins: alternative interactions and oligomeric assemblies. Mol. Vis. 10, 655–662 (2004).
Li, Y., Schmitz, K. R., Salerno, J. C. & Koretz, J. F. The role of the conserved COOH-terminal triad in αA-crystallin aggregation and functionality. Mol. Vis. 13, 1758–1768 (2007).
Alderson, T. R. et al. Local unfolding of the HSP27 monomer regulates chaperone activity. Nat. Commun. 10, 1068 (2019).
Chen, J., Feige, M., Franzmann, T. M., Bepperling, A. & Buchner, J. Regions outside the α-crystallin domain of the small heat shock protein Hsp26 are required for its dimerization. J. Mol. Biol. 398, 122–131 (2010).
Huber-Wunderlich, M. & Glockshuber, R. A single dipeptide sequence modulates the redox properties of a whole enzyme family. Fold Des. 3, 161–171 (1998).
Bova, L. M., Sweeney, M. H., Jamie, J. F. & Truscott, R. J. W. Major changes in human ocular UV protection with age. Invest. Ophthalmol. Vis. Sci. 42, 200–205 (2001).
Grey, A. C., Demarais, N. J., Brandi, J., West, B. J. & Donaldson, P. J. A quantitative map of glutathione in the aging human lens. Int. J. Mass Spectrom. 437, 58–68 (2017).
Hogg, P. J. Disulfide bonds as switches for protein function. Trends Biochem. Sci. 28, 210–214 (2003).
Lou, M. F. Redox regulation in the lens. Prog. Retin. Eye Res. 22, 657–682 (2003).
Rost, J. & Rapoport, S. Reduction-potential of glutathione. Nature 201, 185 (1964).
Simpson, R. J. Estimation of free thiols and disulfide bonds using Ellman’s reagent. Cold Spring Harb. Protoc. 9, 1–8 (2008).
Stafford, W. F. III Boundary analysis in sedimentation transport experiments: a procedure for obtaining sedimentation coefficient distributions using the time derivative of the concentration profile. Anal. Biochem. 203, 295–301 (1992).
Royer, C. A., Mann, C. J. & Matthews, C. R. Resolution of the fluorescence equilibrium unfolding profile of trp aporepressor using single tryptophan mutants. Protein Sci. 11, 1844–1852 (1993).
Wei, H. et al. Using hydrogen/deuterium exchange mass spectrometry to study conformational changes in granulocyte colony stimulating factor upon PEGylation. J. Am. Soc. Mass Spectrom. 23, 498–504 (2012).
Korinek, A., Beck, F., Baumeister, W., Nickell, S. & Plitzko, J. M. Computer controlled cryo-electron microscopy—TOM2 a software package for high-throughput applications. J. Struct. Biol. 175, 394–405 (2011).
Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).
Heymann, J. B. & Belnap, D. M. Bsoft: image processing and molecular modeling for electron microscopy. J. Struct. Biol. 157, 3–18 (2007).
van Heel, M., Harauz, G., Orlova, E. V., Schmidt, R. & Schatz, M. A new generation of the IMAGIC image processing system. J. Struct. Biol. 116, 17–24 (1996).
van Heel, M. Angular reconstitution: a posteriori assignment of projection directions for 3D reconstruction. Ultramicroscopy 21, 111–123 (1987).
Rosenthal, P. B. & Henderson, R. Optimal determination of particle orientation, absolute hand and contrast loss in single-particle electron cryomicroscopy. J. Mol. Biol. 333, 721–745 (2003).
Lawson, C. L. et al. EMDataBank unified data resource for 3DEM. Nucleic Acids Res. 44, D386–D403 (2016).
Goddard, T. D., Huang, C. C. & Ferrin, T. E. Visualizing density maps with UCSF Chimera. J. Struct. Biol. 157, 281–287 (2007).
Penczek, P. A., Yang, C., Frank, J. & Spahn, C. M. Estimation of variance in single-particle reconstruction using the bootstrap technique. J. Struct. Biol. 154, 168–183 (2006).
Maiolica, A. et al. Structural analysis of multiprotein complexes by cross-linking, mass spectrometry and database searching. Mol. Cell. Proteomics 6, 2200–2211 (2007).
Cox, J. & Mann, M. MaxQuant enables high peptide identification rates, individualized p.p.b.-range mass accuracies and proteome-wide protein quantification. Nat. Biotechnol. 26, 1367–1372 (2008).
Mendes, M. L. et al. An integrated workflow for crosslinking mass spectrometry. Mol. Syst. Biol. 15, e8994 (2019).
Fischer, L. & Rappsilber, J. Quirks of error estimation in cross-linking/mass spectrometry. Anal. Chem. 89, 3829–3833 (2017).
Webb, B. & Sali, A. Comparative protein structure modeling using MODELLER. Curr. Protoc. Bioinformatics 47, 1–32 (2014).
Roy, A., Kucukural, A. & Zhang, Y. I-TASSER: a unified platform for automated protein structure and function prediction. Nat. Protoc. 5, 725–738 (2010).
Chacón, P. & Wriggers, W. Multi-resolution contour-based fitting of macromolecular structures. J. Mol. Biol. 317, 375–384 (2002).
Case, D. A. et al. AMBER 16 (University of California, 2016).
Wu, X., Subramaniam, S., Case, D. A., Wu, K. W. & Brooks, B. R. Targeted conformational search with map-restrained self-guided Langevin dynamics: application to flexible fitting into electron microscopic density maps. J. Struct. Biol. 183, 429–440 (2013).
Acknowledgements
We are grateful to J. Plitzko (Max Planck Institute for Biochemistry) for continuous support with EM and critical discussions. We thank D. Balchin for his comments on H/DX-MS data analysis, and M.-L. Jokisch, R. Ciccone and G. Feind for technical assistance during initial experiments. This work was supported by grants from the Deutsche Forschungsgemeinschaft (SFB 1035) and CIPSM to J.B., B. Reif, M.Z. and S.W. Cross-linking/mass spectrometry work was supported by the Wellcome Trust (103139). The Wellcome Centre for Cell Biology is supported by core funding from the Wellcome Trust (203149).
Author information
Authors and Affiliations
Contributions
C.J.O.K., J.B. and S.W. designed and conceived the research plan. C.P., B. Rockel and C.J.O.K. performed EM experiments and processed the data. C.J.O.K. carried out, with contributions from P.W.N.S., E.V.M. and M.H., the experiments for the biochemical and biophysical characterization. V.D. provided full-length human recombinant p53. M.S. and S.A. performed NMR experiments. M.S. and B. Reif analyzed the NMR data. J.Z. conducted cross-linking/mass spectrometry experiments. J.Z. and J.R. analyzed the cross-linking data. M.Z. performed molecular dynamics simulations and model building. C.J.O.K., J.B. and S.W. wrote the manuscript, with input from all authors.
Corresponding authors
Ethics declarations
Competing interests
The authors declare no competing interests.
Additional information
Peer review information Inês Chen was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data
Extended Data Fig. 1 Cryo-EM/3D reconstruction of human αA-crystallin oligomers.
a, Cryo-EM micrograph of human αA-crystallin (reduced, αAred). Top- and side-views are highlighted by white and black circles, respectively. Scale bar: 50 nm. b, Reference-free 2D class averages of top-views (top and middle rows) with corresponding eigenimages indicating size variations and 3-, 4- and 5-fold symmetries (bottom row). c, Reference-free 2D class averages of side-views (top and middle rows) with corresponding eigenimages indicating 2-fold symmetry (bottom row). d–f, Characteristic final class averages (top row) with the corresponding 2D reprojections of the 3D model (bottom row) of the αA-crystallin 12-mer (d), 16-mer (e) and 20-mer (f). Box size in b–f, 17.3 nm. g, Angular distribution plots, that is the distributions of the Euler angles of the final class averages contributing to the 3D reconstructions of the αA-crystallin 12-, 16- and 20-mer. h, Fourier shell correlation (FSC) curves between maps from two independently refined half data sets of 12-, 16- and 20-mer populations. According to the 0.143 gold standard criterion, the resolutions for 12-, 16- and 20-mer 3D reconstructions are 9.2, 9.8 and 9.0 Å, respectively.
Extended Data Fig. 2 Structural variability of human αA-crystallin (reduced) oligomers.
a, c, e, Top and side views of the cryo-EM maps of αA-crystallin (reduced) 12-mer (a), 16-mer (c) and 20-mer (e) (mesh presentation) overlaid with the most important 3D eigenvector (red) indicating the positions of main variances (variance map). b, d, f, Representative 3D class averages of the 12-mer (b), 16-mer (d) and 20-mer (f). The map used for modeling of the 16-mer in 3D domain-swapped configuration is marked in (d) by an asterisk. Scale bar, 10 nm.
Extended Data Fig. 3 Cross-linking of human αA-crystallin.
a, Cross-linker titration of αA-crystallin, denaturing NuPAGE gel. Reduced (left) and oxidized (right) αA-crystallin were incubated for 1 h at room temperature with BS3 cross-linker at the indicated molar BS3:αA-crystallin ratios. Excised monomer (450:1, blue), dimer (450:1 and 900:1, red) and oligomer (450:1, green) gel bands for both αAred and αAox were digested with trypsin and further analyzed. Sequence coverages: αAred-monomer: 97.1 %, αAred-dimer: 99.4 %, αAred-oligomer: 100 %, αAox-monomer: 83.2 %, αAox-dimer: 94.8 %, αAox-oligomer: 100 %. b, Fragmentation spectrum of a cross-linked peptide with an intramolecular link between K70 and K99. c, Fragmentation spectrum of a cross-linked peptide with an intermolecular cross-link between M1 and M1.
Extended Data Fig. 4 Cross-links observed in reduced and oxidized human αA-crystallin.
a, Primary sequence of human αA-crystallin. BS3 reactive K, S, T, Y residues and the N-terminus are coloured red. b, Linkage maps comparing the cross-linked residue pairs observed in monomer, dimer and oligomer pools of αAred and αAox. In total, 113 auto-validation cross-links are shown. Colour code: blue, shared cross-links between αAred and αAox (44 shared cross-links, 39 %); black, unique cross-links in αAox (63 cross-links, 56 %); orange, unique cross-links in αAred (6 cross-links, 5 %). Colour code for the sequence regions of αA-crystallin: NTR (residues 1–60), sienna; ACD (residues 61-145), gray; CTR (residues 146-173), green. c, Histograms of Cα-Cα distances of cross-links observed in αAred. The distances were measured between corresponding residues resolved in the crystal structures of truncated versions of zebrafish (PDB 3N3E, left) and bovine (PDB 3L1E, right) αA-crystallin.
Extended Data Fig. 5 Secondary structure prediction and modeling of the N-terminal region of human αA-crystallin.
a, Summary of sequence-based secondary structure predictions of the NTR as obtained from 15 different web-based prediction programs. The predictions reproduce all β-strand segments (blue) present in metazoan sHsp structures. According to the predictions, the NTR most likely contains 3-4 α-helical segments (orange). b, A possible 3D structure model of the NTR of human αA-crystallin predicted using I-Tasser. c, Examples of possible conformations of the NTR of apical (Map) and d, equatorial protomers (Meq) obtained upon structure modeling by molecular dynamics flexible fitting. Although the positions of the three helices within the EM-density in both Map and Meq differ, their arrangement relative to each other is well preserved in comparison to the I-Tasser model mRMSD ∼2 Å).
Extended Data Fig. 6 Superposition of 1H,15N correlation spectra of 15N-αAred and 15N-αAred-IPSL.
The superposition of 1H,15N correlation spectra of 15N-αAred (black) and 15N-αAred-IPSL treated with ascorbic acid (reduced) shows chemical shift perturbations for residues, for which we have observed an attenuation of the signal intensity for the oxidized 15N-αAred -IPSL sample. In particular, residues T153, A155, E156, R157 display significant chemical shift changes, consistent with the PRE results. At the same time, the chemical shifts of the C-terminal residues (T168, S169, A170, S172, S173) are not affected by the presence of the nitroxyl moiety.
Extended Data Fig. 7 Impact of oxidation on αA-crystallin structure.
a,b, Far-UV (a) and near-UV (b) CD spectra of αAred (black line) and αAox (gray line). Note that the chemical microenvironment of tyrosins, phenylalanines and W9 are affected by oxidation. c, SEC elution profiles of αAred (black line) and αAox (gray line) on a Superose 6 10/300 GL column. Inset: a segment of the calibration curve using the filtration standard mixture from BioRad. The calculated average molecular masses are 380 kDa for αAred and 770 kDa for αAox, respectively (ThG: bovine thyroglobulin, 670 kDa; γG: bovine γ-globulin, 158 kDa). Note the peak broadening, that is increased polydispersity in αAox. d, Analysis of αAred (black line) and αAox (gray line) by sedimentation velocity aUC in a concentration range from 2µM to 150 µM using SEDFIT. The concentrations are 2 µM, 10 µM, 20 µM, 50 µM and 150 µM. The inset shows the concentration dependence of the sedimentation coefficient. e, A set of the class averages used for the 3D reconstruction of αAox 32-mer. f, 2D reprojections of the reconstructed 3D volume corresponding to the orientations of the class averages shown in (e). Box size in e and f: 26.7 nm.
Extended Data Fig. 8 Impact of oxidation on αA-crystallin stability.
a, Oligomeric states of αAred (black circles) and αAox (gray circles) in the presence of urea as determined by sedimentation velocity aUC at 20 °C. The oligomers of both proteins dissociate successively with increasing urea concentrations. Note that αAred and αAox form a ∼2S species at urea concentrations of 4.5 M and 3.5 M, respectively, suggesting destabilization of αAox oligomers. b, Intrinsic fluorescence urea unfolding transitions for αAred and αAox at 20 °C. The midpoints of the cooperative transition are at 2.7 M for αAox and at 3.8 M urea for αAred, indicating destabilization of the NTR in the case of αAox. The spectral settings of the fluorimeter were chosen to selectively assess the transition of W9 located within the NTR.
Extended Data Fig. 9 αA-crystallin is capable of transferring disulfide bonds to MDH.
a, Denaturing, non-reducing PAGE of samples withdrawn at the indicated timepoints (red arrows) from the aggregation assays in the presence of αAred and recombinant reduced E. coli DsbA as shown in Fig. 7a. Note that disulfide-bridged species of p53 are formed only in marginal amounts. b, Heat-induced aggregation of recombinant malate dehydrogenase (MDH, 4µM) in the presence of an equimolar amount of GSSG, αAred, αAox and reduced (DsbAred) or oxidized (DsbAox) E. coli DsbA. Note that the aggregation of MDH is fully suppressed in the presence of αAred and almost fully suppressed in the presence of αAox. c, Relative intensity of the MDH monomer band as a fraction of the initial intensity (amount of monomer) at the beginning of each aggregation kinetics experiment (t = 0 min). d,e, Denaturing, non-reducing PAGE of samples withdrawn at the indicated timepoints (red arrows) from the aggregation assays shown in (b). Experiments were performed in the presence of GSSG, αAox or DsbAox (d), in the absence of GSSG (MDH only) and in the presence of αAred or DsbAred (e). Note that disulfide-bridged species of MDH are formed in the presence of αAox.
Supplementary information
Supplementary Information
Supplementary Note, Supplementary Fig. 1 and Table 1.
Supplementary Table 1
Cross-links observed in monomers, dimers and oligomers of reduced αA-crystallin.
Supplementary Table 2
Cross-links observed in monomers, dimers and oligomers of oxidized αA-crystallin.
Rights and permissions
About this article
Cite this article
Kaiser, C.J.O., Peters, C., Schmid, P.W.N. et al. The structure and oxidation of the eye lens chaperone αA-crystallin. Nat Struct Mol Biol 26, 1141–1150 (2019). https://doi.org/10.1038/s41594-019-0332-9
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41594-019-0332-9
This article is cited by
-
An insight into the structural analysis of α-crystallin of habitat-specific fish: a computational approach
Journal of Proteins and Proteomics (2023)
-
Phosphorylation activates the yeast small heat shock protein Hsp26 by weakening domain contacts in the oligomer ensemble
Nature Communications (2021)
-
Peeking from behind the veil of enigma: emerging insights on small heat shock protein structure and function
Cell Stress and Chaperones (2020)