Structural insights into photoactivation and signalling in plant phytochromes.
Journal
Nature plants
ISSN: 2055-0278
Titre abrégé: Nat Plants
Pays: England
ID NLM: 101651677
Informations de publication
Date de publication:
05 2020
05 2020
Historique:
received:
17
12
2019
accepted:
16
03
2020
pubmed:
6
5
2020
medline:
10
2
2021
entrez:
6
5
2020
Statut:
ppublish
Résumé
Plant phytochromes are red/far-red photochromic photoreceptors that act as master regulators of development, controlling the expression of thousands of genes. Here, we describe the crystal structures of four plant phytochrome sensory modules, three at about 2 Å resolution or better, including the first of an A-type phytochrome. Together with extensive spectral data, these structures provide detailed insight into the structure and function of plant phytochromes. In the Pr state, the substitution of phycocyanobilin and phytochromobilin cofactors has no structural effect, nor does the amino-terminal extension play a significant functional role. Our data suggest that the chromophore propionates and especially the phytochrome-specific domain tongue act differently in plant and prokaryotic phytochromes. We find that the photoproduct in period-ARNT-single-minded (PAS)-cGMP-specific phosphodiesterase-adenylyl cyclase-FhlA (GAF) bidomains might represent a novel intermediate between MetaRc and Pfr. We also discuss the possible role of a likely nuclear localization signal specific to and conserved in the phytochrome A lineage.
Identifiants
pubmed: 32366982
doi: 10.1038/s41477-020-0638-y
pii: 10.1038/s41477-020-0638-y
doi:
Substances chimiques
Phytochrome
11121-56-5
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
581-588Références
Anders, K., Daminelli-Widany, G., Mroginski, M. A., von Stetten, D. & Essen, L. O. Structure of the cyanobacterial phytochrome 2 photosensor implies a tryptophan switch for phytochrome signaling. J. Biol. Chem. 288, 35714–35725 (2013).
pubmed: 24174528
pmcid: 3861623
Takala, H. et al. Signal amplification and transduction in phytochrome photosensors. Nature 509, 245–248 (2014).
pubmed: 24776794
pmcid: 4015848
Oka, Y. et al. Functional analysis of a 450-amino acid N-terminal fragment of phytochrome B in Arabidopsis. Plant Cell 16, 2104–2116 (2004).
pubmed: 15273294
pmcid: 519201
Matsushita, T., Mochizuki, N. & Nagatani, A. Dimers of the N-terminal domain of phytochrome B are functional in the nucleus. Nature 424, 571–574 (2003).
pubmed: 12891362
Qiu, Y. et al. Mechanism of early light signaling by the carboxy-terminal output module of Arabidopsis phytochrome B. Nat. Commun. 8, 1905 (2017).
pubmed: 29199270
pmcid: 5712524
Viczian, A. et al. A short amino-terminal part of Arabidopsis phytochrome A induces constitutive photomorphogenic response. Mol. Plant 5, 629–641 (2012).
pubmed: 22498774
Song, C. et al. 3D structures of plant phytochrome A as Pr and Pfr from solid-state NMR: implications for molecular function. Front. Plant Sci. 9, 498 (2018).
pubmed: 29740459
pmcid: 5928327
Sharrock, R. A. & Quail, P. H. Novel phytochrome sequences in Arabidopsis thaliana: structure, evolution, and differential expression of a plant regulatory photoreceptor family. Genes Dev. 3, 1745–1757 (1989).
pubmed: 2606345
Rockwell, N. C., Shang, L., Martin, S. S. & Lagarias, J. C. Distinct classes of red/far-red photochemistry within the phytochrome superfamily. Proc. Natl Acad. Sci. USA 106, 6123–6127 (2009).
pubmed: 19339496
Mailliet, J. et al. Spectroscopy and a high-resolution crystal structure of Tyr263 mutants of cyanobacterial phytochrome Cph1. J. Mol. Biol. 413, 115–127 (2011).
pubmed: 21888915
Song, C. et al. Two ground state isoforms and a chromophore D-ring photoflip triggering extensive intramolecular changes in a canonical phytochrome. Proc. Natl Acad. Sci. USA 108, 3842–3847 (2011).
pubmed: 21325055
Essen, L.-O., Mailliet, J. & Hughes, J. The structure of a complete phytochrome sensory module in the Pr ground state. Proc. Natl Acad. Sci. USA 105, 14709–14714 (2008).
pubmed: 18799745
Burgie, E. S., Bussell, A. N., Walker, J. M., Dubiel, K. & Vierstra, R. D. Crystal structure of the photosensing module from a red/far-red light-absorbing plant phytochrome. Proc. Natl Acad. Sci. USA 111, 10179–10184 (2014).
pubmed: 24982198
Kneip, C. et al. Effect of chromophore exchange on the resonance Raman spectra of recombinant phytochromes. FEBS 414, 23–26 (1997).
Remberg, A. et al. Raman spectroscopic and light-induced-kinetic characterization of a recombinant phytochrome of the cyanobacterium Synechocystis. Biochemistry 36, 13389–13395 (1997).
pubmed: 9341232
Mroginski, M. A. et al. Structure of the chromophore binding pocket in the Pr state of plant phytochrome phyA. J. Phys. Chem. B 115, 1220–1231 (2011).
pubmed: 21192668
Wahleithner, J. A., Li, L. M. & Lagarias, J. C. Expression and assembly of spectrally active recombinant holophytochrome. Proc. Natl Acad. Sci. USA 88, 10387–10391 (1991).
pubmed: 1961704
Elich, T. D. & Lagarias, J. C. Formation of a photoreversible phycocyanobilin-apophytochrome adduct in vitro. J. Biol. Chem. 264, 12902–12908 (1989).
pubmed: 2753895
Song, C., Essen, L. O., Gärtner, W., Hughes, J. & Matysik, J. Solid-state NMR spectroscopic study of chromophore–protein interactions in the Pr ground state of plant phytochrome A. Mol. Plant 5, 698–715 (2012).
pubmed: 22419823
Velazquez Escobar, F. et al. Structural communication between the chromophore-binding pocket and the N-terminal extension in plant phytochrome phyB. FEBS Lett. 591, 1258–1265 (2017).
pubmed: 28376244
Parker, W., Partis, M. & Song, P. S. N-terminal domain of Avena phytochrome: interactions with sodium dodecyl sulfate micelles and N-terminal chain truncated phytochrome. Biochemistry 31, 9413–9420 (1992).
pubmed: 1390724
Litts, J. C., Kelly, J. M. & Lagarias, J. C. Structure–function studies on phytochrome: preliminary characterization of highly purified phytochrome from Avena sativa enriched in the 124-kilodalton species. J. Biol. Chem. 258, 11025–11031 (1983).
pubmed: 6885811
Claesson, E. et al. The primary structural photoresponse of phytochrome proteins captured by a femtosecond X-ray laser. eLife 9, e53514 (2020).
pubmed: 32228856
pmcid: 7164956
Wagner, J. R., Brunzelle, J. S., Forest, K. T. & Vierstra, R. D. A light-sensing knot revealed by the structure of the chromophore-binding domain of phytochrome. Nature 438, 325–331 (2005).
pubmed: 16292304
Eilfeld, P. & Rüdiger, W. Absorption spectra of phytochrome intermediates. Z. Naturforsch. 40c, 109–114 (1985).
Kneip, C. et al. Protonation state and structural changes of the tetrapyrrole chromophore during the Pr → Pfr phototransformation of phytochrome: a resonance Raman spectroscopic study. Biochemistry 38, 15185–15192 (1999).
pubmed: 10563801
Mizutani, Y., Tokutomi, S. & Kitagawa, T. Resonance Raman spectra of the intermediates in phototransformation of large phytochrome: deprotonation of the chromophore in the bleached intermediate. Biochemistry 33, 153–158 (1994).
pubmed: 8286333
Mroginski, M. A. et al. Elucidating photoinduced structural changes in phytochromes by the combined application of resonance Raman spectroscopy and theoretical methods. J. Mol. Struct. 993, 15–25 (2011).
Ulijasz, A. T. et al. Characterization of two thermostable cyanobacterial phytochromes reveals global movements in the chromophore-binding domain during photoconversion. J. Biol. Chem. 283, 21251–21266 (2008).
pubmed: 18480055
pmcid: 3258942
Song, C. et al. The D-ring, not the A-ring, rotates in Synechococcus OS-B’ phytochrome. J. Biol. Chem. 289, 2552–2562 (2014).
pubmed: 24327657
Auldridge, M. E., Satyshur, K. A., Anstrom, D. M. & Forest, K. T. Structure-guided engineering enhances a phytochrome-based infrared fluorescent protein. J. Biol. Chem. 287, 7000–7009 (2012).
pubmed: 22210774
Ihalainen, J. A. et al. Chromophore–protein interplay during the phytochrome photocycle revealed by step-scan FTIR spectroscopy. J. Am. Chem. Soc. 140, 12396–12404 (2018).
pubmed: 30183281
Kacprzak, S. et al. Intersubunit distances in full-length, dimeric, bacterial phytochrome Agp1, as measured by pulsed electron-electron double resonance (PELDOR) between different spin label positions, remain unchanged upon photoconversion. J. Biol. Chem. 292, 7598–7606 (2017).
pubmed: 28289094
pmcid: 5418057
Park, E. et al. Phytochrome B inhibits binding of phytochrome-interacting factors to their target promoters. Plant J. 72, 537–546 (2012).
pubmed: 22849408
pmcid: 3489987
Oka, Y., Matsushita, T., Mochizuki, N., Quail, P. H. & Nagatani, A. Mutant screen distinguishes between residues necessary for light-signal perception and signal transfer by phytochrome B. PLoS Genet. 4, e1000158 (2008).
pubmed: 18704165
pmcid: 2494609
Kikis, E. A., Oka, Y., Hudson, M. E., Nagatani, A. & Quail, P. H. Residues clustered in the light-sensing knot of phytochrome B are necessary for conformer-specific binding to signaling partner PIF3. PLoS Genet. 5, e1000352 (2009).
pubmed: 19165330
pmcid: 2621353
Viczian, A., Klose, C., Adam, E. & Nagy, F. New insights of red light-induced development. Plant Cell Environ. 40, 2457–2468 (2017).
pubmed: 27943362
Burgie, E. S., Zhang, J. & Vierstra, R. D. Crystal structure of Deinococcus phytochrome in the photoactivated state reveals a cascade of structural rearrangements during photoconversion. Structure 24, 448–457 (2016).
pubmed: 26853942
Yang, X., Kuk, J. & Moffat, K. Conformational differences between the Pfr and Pr states in Pseudomonas aeruginosa bacteriophytochrome. Proc. Natl Acad. Sci. USA 106, 15639–15644 (2009).
pubmed: 19720999
van Thor, J. J. et al. Light-induced proton release and proton uptake reactions in the cyanobacterial phytochrome Cph1. Biochemistry 40, 11460–11471 (2001).
pubmed: 11560494
Borucki, B. et al. Light-induced proton release of phytochrome is coupled to the transient deprotonation of the tetrapyrrole chromophore. J. Biol. Chem. 280, 34358–34364 (2005).
pubmed: 16061486
Viczian, A. et al. Differential phosphorylation of the N-terminal extension regulates phytochrome B signaling. New Phytol. 225, 1635–1650 (2020).
pubmed: 31596952
Remberg, A., Ruddat, A., Braslavsky, S. E., Gärtner, W. & Schaffner, K. Chromophore incorporation, Pr to Pfr kinetics, and Pfr thermal reversion of recombinant N-terminal fragments of phytochrome A and B chromoproteins. Biochemistry 37, 9983–9990 (1998).
pubmed: 9665703
Cherry, J. R., Hondred, D., Walker, J. M. & Vierstra, R. D. Phytochrome requires the 6-kDa N-terminal domain for full biological activity. Proc. Natl Acad. Sci. USA 89, 5039–5043 (1992).
pubmed: 1594611
Sweere, U. et al. Interaction of the response regulator ARR4 with phytochrome B in modulating red light signaling. Science 294, 1108–1111 (2001).
pubmed: 11691995
Kosugi, S. et al. Six classes of nuclear localization signals specific to different binding grooves of importin alpha. J. Biol. Chem. 284, 478–485 (2009).
pubmed: 19001369
Mathews, S., Lavin, M. & Sharrock, R. A. Evolution of the phytochrome gene family and its utility for phylogenetic analyses of angiosperms. Ann. Missouri Bot. Gard. 82, 296–321 (1995).
Chen, M. & Chory, J. Phytochrome signaling mechanisms and the control of plant development. Trends Cell Biol. 21, 664–671 (2011).
pubmed: 21852137
pmcid: 3205231
Zeidler, M., Zhou, Q., Sarda, X., Yau, C. P. & Chua, N. H. The nuclear localization signal and the C-terminal region of FHY1 are required for transmission of phytochrome A signals. Plant J. 40, 355–365 (2004).
pubmed: 15469493
Oka, Y. et al. Arabidopsis phytochrome A is modularly structured to integrate the multiple features that are required for a highly sensitized phytochrome. Plant Cell 24, 2949–2962 (2012).
pubmed: 22843485
pmcid: 3426125
Song, C. et al. On the collective nature of phytochrome photoactivation. Biochemistry 50, 10987–10989 (2011).
pubmed: 22124256
Kim, P. W. et al. Unraveling the primary isomerization dynamics in cyanobacterial phytochrome Cph1 with multi-pulse manipulations. J. Phys. Chem. Lett. 4, 2605–2609 (2013).
pubmed: 24143267
pmcid: 3798021
Gustavsson, E. et al. Modulation of structural heterogeneity controls phytochrome photoswitching. Biophys. J. 118, 415–421 (2020).
pubmed: 31839260
Takala, H. et al. On the (un)coupling of the chromophore, tongue interactions, and overall conformation in a bacterial phytochrome. J. Biol. Chem. 293, 8161–8172 (2018).
pubmed: 29622676
pmcid: 5971452
Al-Sady, B., Ni, W. M., Kircher, S., Schäfer, E. & Quail, P. H. Photoactivated phytochrome induces rapid PIF3 phosphorylation prior to proteasorne-mediated degradation. Mol. Cell 23, 439–446 (2006).
pubmed: 16885032
Khanna, R. et al. A novel molecular recognition motif necessary for targeting photoactivated phytochrome signaling to specific basic helix-loop-helix transcription factors. Plant Cell 16, 3033–3044 (2004).
pubmed: 15486100
pmcid: 527196
Landgraf, F. T., Forreiter, C., Hurtado, P. A., Lamparter, T. & Hughes, J. Recombinant holophytochrome in Escherichia coli. FEBS Lett. 508, 459–462 (2001).
pubmed: 11728472
Hirose, Y. et al. Green/red cyanobacteriochromes regulate complementary chromatic acclimation via a protochromic photocycle. Proc. Natl Acad. Sci. USA 110, 4974–4979 (2013).
pubmed: 23479641
Zhao, K. H. & Scheer, H. Type I and type II reversible photochemistry of phycoerythrocyanin α-subunit from Mastigocladus laminosus both involve Z–E isomerization of phycoviolobilin chromophore and are controlled by sulfhydryls in apoprotein. Biochim. Biophys. Acta Bioenerg. 1228, 244–253 (1995).
Ishizuka, T., Narikawa, R., Kohchi, T., Katayama, M. & Ikeuchi, M. Cyanobacteriochrome TePixJ of Thermosynechococcus elongatus harbors phycoviolobilin as a chromophore. Plant Cell Physiol. 48, 1385–1390 (2007).
pubmed: 17715149
Brandlmeier, T., Scheer, H. & Rudiger, W. Chromophore content and molar absorptivity of phytochrome in the Pr form. Z. Naturforsch. C 36, 431–439 (1981).
Fernandez Lopez, M. et al. Role of the propionic side chains for the photoconversion of bacterial phytochromes. Biochemistry 58, 3504–3519 (2019).
pubmed: 31348653
Mailliet, J. et al. Dwelling in the dark: procedures for the crystallography of phytochromes and other photochromic proteins. Acta Crystallogr. D 65, 1232–1235 (2009).
pubmed: 19923720
Kabsch, W. XDS. Acta Crystallogr. D 66, 125–132 (2010).
pubmed: 20124692
Evans, P. R. & Murshudov, G. N. How good are my data and what is the resolution? Acta Crystallogr. D 69, 1204–1214 (2013).
pubmed: 23793146
Winn, M. D. et al. Overview of the CCP4 suite and current developments. Acta Crystallogr. D 67, 235–242 (2011).
pubmed: 21460441
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007).
pubmed: 19461840
pmcid: 19461840
Emsley, P. B., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D 66, 486–501 (2010).
pubmed: 20383002
Murshudov, G. N. et al. REFMAC5 for the refinement of macromolecular crystal structures. Acta Crystallogr. D 67, 355–367 (2011).
pubmed: 21460454
Liebschner, D. et al. Macromolecular structure determination using X-rays, neutrons and electrons: recent developments in Phenix. Acta Crystallogr. D 75, 861–877 (2019).
Williams, C. J. et al. MolProbity: more and better reference data for improved all-atom structure validation. Protein Sci. 27, 293–315 (2018).
pubmed: 29067766
Dolomanov, O. V., Bourhis, L. J., Gildea, R. J., Howard, J. A. K. & Puschmann, H. OLEX2: a complete structure solution, refinement and analysis program. J. Appl. Crystallogr. 42, 339–341 (2009).