The structure and oxidation of the eye lens chaperone αA-crystallin.
Journal
Nature structural & molecular biology
ISSN: 1545-9985
Titre abrégé: Nat Struct Mol Biol
Pays: United States
ID NLM: 101186374
Informations de publication
Date de publication:
12 2019
12 2019
Historique:
received:
24
04
2019
accepted:
10
10
2019
pubmed:
4
12
2019
medline:
26
2
2020
entrez:
4
12
2019
Statut:
ppublish
Résumé
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.
Identifiants
pubmed: 31792453
doi: 10.1038/s41594-019-0332-9
pii: 10.1038/s41594-019-0332-9
pmc: PMC7115824
mid: EMS84590
doi:
Substances chimiques
alpha-Crystallin A Chain
0
Types de publication
Journal Article
Research Support, Non-U.S. Gov't
Langues
eng
Sous-ensembles de citation
IM
Pagination
1141-1150Subventions
Organisme : Wellcome Trust
Pays : United Kingdom
Organisme : Wellcome Trust
ID : 103139
Pays : United Kingdom
Organisme : Wellcome Trust
ID : 103139/Z/13/Z
Pays : United Kingdom
Organisme : Wellcome Trust
ID : 203149
Pays : United Kingdom
Références
Bloemendal, H. et al. Ageing and vision: structure, stability and function of lens crystallins. Prog. Biophys. Mol. Biol. 86, 407–485 (2004).
pubmed: 15302206
doi: 10.1016/j.pbiomolbio.2003.11.012
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).
pubmed: 22405853
doi: 10.1016/j.biocel.2012.02.015
Horwitz, J. α-Crystallin can function as a molecular chaperone. Proc. Natl Acad. Sci. USA 89, 10449–10453 (1992).
pubmed: 1438232
pmcid: 50356
doi: 10.1073/pnas.89.21.10449
Horwitz J. Alpha-crystallin. Exp. Eye Res. 76, 145–153 (2003).
pubmed: 12565801
doi: 10.1016/S0014-4835(02)00278-6
Graw, J. Genetics of crystallins: cataract and beyond. Exp. Eye Res. 88, 173–189 (2009).
pubmed: 19007775
doi: 10.1016/j.exer.2008.10.011
Oguni, M. et al. Ontogeny of alpha-crystallin subunits in the lens of human and rat embryos. Cell Tissue Res. 276, 151–154 (1994).
pubmed: 8187157
doi: 10.1007/BF00354794
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).
pubmed: 2539261
doi: 10.1016/0092-8674(89)90173-6
Srinivasan, A. N., Nagineni, C. N. & Bhat, S. P. αA-crystallin is expressed in non-ocular tissues. J. Biol. Chem. 267, 23337–23341 (1992).
pubmed: 1429679
doi: 10.1016/S0021-9258(18)50096-X
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).
pubmed: 23071119
pmcid: 3516784
doi: 10.1074/jbc.M112.414854
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).
pubmed: 10574947
doi: 10.1074/jbc.274.49.34773
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).
pubmed: 10671481
doi: 10.1074/jbc.275.7.4565
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).
pubmed: 10446186
doi: 10.1074/jbc.274.34.24137
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).
pubmed: 7723051
doi: 10.1007/BF00163229
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).
pubmed: 9650070
doi: 10.1016/S0141-8130(98)00013-0
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).
pubmed: 20440841
pmcid: 2868245
doi: 10.1002/pro.380
Laganowsky, A. & Eisenberg, D. Non-3D domain swapped crystal structure of truncated zebrafish alphaA crystallin. Protein Sci. 19, 1978–1984 (2010).
pubmed: 20669149
pmcid: 2998731
doi: 10.1002/pro.471
Bova, M. P., Ding, L. L., Horwitz, J. & Fung, B. K. Subunit exchange of αA-crystallin. J. Biol. Chem. 272, 29511–29517 (1997).
pubmed: 9368012
doi: 10.1074/jbc.272.47.29511
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).
pubmed: 15701626
doi: 10.1074/jbc.M500135200
Peschek, J. et al. The eye lens chaperone α-crystallin forms defined globular assemblies. Proc. Natl Acad. Sci. USA 106, 13272–13277 (2009).
pubmed: 19651604
pmcid: 2726422
doi: 10.1073/pnas.0902651106
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).
pubmed: 1562604
doi: 10.1016/0167-4781(92)90439-7
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).
pubmed: 10625643
doi: 10.1074/jbc.275.2.1035
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).
pubmed: 14631074
doi: 10.1093/protein/gzg102
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).
pubmed: 17345631
doi: 10.1002/bip.20716
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).
pubmed: 8943244
doi: 10.1074/jbc.271.50.31973
Thampi, P. & Abraham, E. C. Influence of the C-terminal residues on oligomerization of αA-crystallin. Biochemistry 42, 11857–11863 (2003).
pubmed: 14529298
doi: 10.1021/bi030129w
Rajan, S., Chandrashekar, R., Aziz, A. & Abraham, E. C. Role of arginine-163 and the
pubmed: 17176090
doi: 10.1021/bi060705z
Aziz, A., Santhoshkumar, P., Sharma, K. K. & Abraham, E. C. Cleavage of the C-terminal serine of human αA-crystallin produces αA
pubmed: 17279772
doi: 10.1021/bi0618722
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).
pubmed: 20732317
doi: 10.1016/j.exer.2010.08.015
Kim, K. K., Kim, R. & Kim, S. H. Crystal structure of a small heat-shock protein. Nature 394, 595–599 (1998).
pubmed: 9707123
doi: 10.1038/29106
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).
pubmed: 11702068
doi: 10.1038/nsb722
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).
pubmed: 11925526
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).
pubmed: 3620480
doi: 10.1016/0167-4838(87)90133-6
Srikanthan, D., Bateman, O. A., Purkiss, A. G. & Slingsby, C. Sulfur in human crystallins. Exp. Eye Res. 79, 823–831 (2004).
pubmed: 15642319
doi: 10.1016/j.exer.2004.05.009
Miesbauer, L. R. et al. Post-translational modifications of water-soluble human lens crystallins from young adult. J. Biol. Chem. 269, 12494–12502 (1994).
pubmed: 8175657
doi: 10.1016/S0021-9258(18)99902-3
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).
pubmed: 8670261
doi: 10.1006/bbrc.1996.0873
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).
pubmed: 8994362
doi: 10.1006/exer.1996.0149
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).
pubmed: 18761110
doi: 10.1016/j.bbapap.2008.07.016
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).
pubmed: 26453637
pmcid: 4762625
doi: 10.1074/mcp.M115.050948
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).
pubmed: 7924405
doi: 10.3109/02713689408999869
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).
pubmed: 9068373
doi: 10.1006/exer.1996.0160
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).
pubmed: 10930324
doi: 10.1006/exer.2000.0868
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).
pubmed: 10544964
doi: 10.1023/A:1006906615469
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).
pubmed: 24639379
pmcid: 4093951
doi: 10.1002/pro.2458
Dyson, H. J. & Wright, P. E. Unfolded proteins and protein folding studied by NMR. Chem. Rev. 104, 3607–3622 (2004).
pubmed: 15303830
doi: 10.1021/cr030403s
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).
pubmed: 19522502
pmcid: 2825090
doi: 10.1021/cr900033p
Chakraborty, K. et al. Protein stabilization by introduction of cross-strand disulfides. Biochemistry 44, 14638–14646 (2005).
pubmed: 16262263
doi: 10.1021/bi050921s
Wunderlich, M. & Glockshuber, R. Redox properties of protein disulfide isomerase (DsbA) from Escherichia coli. Protein Sci. 2, 717–726 (1993).
pubmed: 8495194
pmcid: 2142495
doi: 10.1002/pro.5560020503
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).
pubmed: 7536035
doi: 10.1021/bi00015a019
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).
pubmed: 14739319
pmcid: 2286712
doi: 10.1110/ps.03180004
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).
pubmed: 21464278
pmcid: 3081008
doi: 10.1073/pnas.1014656108
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).
pubmed: 26458046
doi: 10.1038/nsmb.3108
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).
pubmed: 28089448
pmcid: 5321513
doi: 10.1016/j.str.2016.12.005
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).
pubmed: 14532291
doi: 10.1074/jbc.M307523200
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).
pubmed: 21839749
doi: 10.1016/j.jmb.2011.07.017
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).
pubmed: 28547731
pmcid: 5465039
doi: 10.1007/s12192-017-0791-z
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).
pubmed: 20802487
pmcid: 2957905
doi: 10.1038/nsmb.1891
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).
pubmed: 22143763
pmcid: 3251151
doi: 10.1073/pnas.1111014108
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).
pubmed: 15448619
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).
pubmed: 17960114
Alderson, T. R. et al. Local unfolding of the HSP27 monomer regulates chaperone activity. Nat. Commun. 10, 1068 (2019).
pubmed: 30842409
pmcid: 6403371
doi: 10.1038/s41467-019-08557-8
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).
pubmed: 20171228
doi: 10.1016/j.jmb.2010.02.022
Huber-Wunderlich, M. & Glockshuber, R. A single dipeptide sequence modulates the redox properties of a whole enzyme family. Fold Des. 3, 161–171 (1998).
pubmed: 9562546
doi: 10.1016/S1359-0278(98)00024-8
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).
pubmed: 11133868
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).
doi: 10.1016/j.ijms.2017.10.008
Hogg, P. J. Disulfide bonds as switches for protein function. Trends Biochem. Sci. 28, 210–214 (2003).
pubmed: 12713905
doi: 10.1016/S0968-0004(03)00057-4
Lou, M. F. Redox regulation in the lens. Prog. Retin. Eye Res. 22, 657–682 (2003).
pubmed: 12892645
doi: 10.1016/S1350-9462(03)00050-8
Rost, J. & Rapoport, S. Reduction-potential of glutathione. Nature 201, 185 (1964).
pubmed: 14118271
doi: 10.1038/201185a0
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).
pubmed: 1416025
doi: 10.1016/0003-2697(92)90316-Y
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).
doi: 10.1002/pro.5560021106
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).
pubmed: 22227798
pmcid: 3438904
doi: 10.1007/s13361-011-0310-x
Korinek, A., Beck, F., Baumeister, W., Nickell, S. & Plitzko, J. M. Computer controlled cryo-electron microscopy—TOM
pubmed: 21704708
doi: 10.1016/j.jsb.2011.06.003
Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).
pubmed: 16859925
doi: 10.1016/j.jsb.2006.05.009
Heymann, J. B. & Belnap, D. M. Bsoft: image processing and molecular modeling for electron microscopy. J. Struct. Biol. 157, 3–18 (2007).
pubmed: 17011211
doi: 10.1016/j.jsb.2006.06.006
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).
pubmed: 8742718
doi: 10.1006/jsbi.1996.0004
van Heel, M. Angular reconstitution: a posteriori assignment of projection directions for 3D reconstruction. Ultramicroscopy 21, 111–123 (1987).
pubmed: 12425301
doi: 10.1016/0304-3991(87)90078-7
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).
pubmed: 14568533
doi: 10.1016/j.jmb.2003.07.013
Lawson, C. L. et al. EMDataBank unified data resource for 3DEM. Nucleic Acids Res. 44, D386–D403 (2016).
doi: 10.1093/nar/gkv1126
Goddard, T. D., Huang, C. C. & Ferrin, T. E. Visualizing density maps with UCSF Chimera. J. Struct. Biol. 157, 281–287 (2007).
pubmed: 16963278
doi: 10.1016/j.jsb.2006.06.010
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).
pubmed: 16510296
doi: 10.1016/j.jsb.2006.01.003
Maiolica, A. et al. Structural analysis of multiprotein complexes by cross-linking, mass spectrometry and database searching. Mol. Cell. Proteomics 6, 2200–2211 (2007).
pubmed: 17921176
doi: 10.1074/mcp.M700274-MCP200
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).
pubmed: 19029910
doi: 10.1038/nbt.1511
Mendes, M. L. et al. An integrated workflow for crosslinking mass spectrometry. Mol. Syst. Biol. 15, e8994 (2019).
pubmed: 31556486
pmcid: 6753376
doi: 10.15252/msb.20198994
Fischer, L. & Rappsilber, J. Quirks of error estimation in cross-linking/mass spectrometry. Anal. Chem. 89, 3829–3833 (2017).
pubmed: 28267312
pmcid: 5423704
doi: 10.1021/acs.analchem.6b03745
Webb, B. & Sali, A. Comparative protein structure modeling using MODELLER. Curr. Protoc. Bioinformatics 47, 1–32 (2014).
doi: 10.1002/0471250953.bi0506s47
Roy, A., Kucukural, A. & Zhang, Y. I-TASSER: a unified platform for automated protein structure and function prediction. Nat. Protoc. 5, 725–738 (2010).
pubmed: 20360767
pmcid: 2849174
doi: 10.1038/nprot.2010.5
Chacón, P. & Wriggers, W. Multi-resolution contour-based fitting of macromolecular structures. J. Mol. Biol. 317, 375–384 (2002).
pubmed: 11922671
doi: 10.1006/jmbi.2002.5438
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).
pubmed: 23876978
pmcid: 3785014
doi: 10.1016/j.jsb.2013.07.006