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.
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The three-dimensional structural data that support the findings of this study have been deposited in wwPDB with the accession codes 6TBY, 6TC5, 6TL4 and 6TC7. The authors declare that all other data supporting the findings of this study are available within the paper and its Supplementary Information files.
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).
Takala, H. et al. Signal amplification and transduction in phytochrome photosensors. Nature 509, 245–248 (2014).
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).
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).
Qiu, Y. et al. Mechanism of early light signaling by the carboxy-terminal output module of Arabidopsis phytochrome B. Nat. Commun. 8, 1905 (2017).
Viczian, A. et al. A short amino-terminal part of Arabidopsis phytochrome A induces constitutive photomorphogenic response. Mol. Plant 5, 629–641 (2012).
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).
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).
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).
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).
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).
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).
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).
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).
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).
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).
Elich, T. D. & Lagarias, J. C. Formation of a photoreversible phycocyanobilin-apophytochrome adduct in vitro. J. Biol. Chem. 264, 12902–12908 (1989).
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).
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).
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).
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).
Claesson, E. et al. The primary structural photoresponse of phytochrome proteins captured by a femtosecond X-ray laser. eLife 9, e53514 (2020).
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).
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).
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).
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).
Song, C. et al. The D-ring, not the A-ring, rotates in Synechococcus OS-B’ phytochrome. J. Biol. Chem. 289, 2552–2562 (2014).
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).
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).
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).
Park, E. et al. Phytochrome B inhibits binding of phytochrome-interacting factors to their target promoters. Plant J. 72, 537–546 (2012).
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).
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).
Viczian, A., Klose, C., Adam, E. & Nagy, F. New insights of red light-induced development. Plant Cell Environ. 40, 2457–2468 (2017).
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).
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).
van Thor, J. J. et al. Light-induced proton release and proton uptake reactions in the cyanobacterial phytochrome Cph1. Biochemistry 40, 11460–11471 (2001).
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).
Viczian, A. et al. Differential phosphorylation of the N-terminal extension regulates phytochrome B signaling. New Phytol. 225, 1635–1650 (2020).
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).
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).
Sweere, U. et al. Interaction of the response regulator ARR4 with phytochrome B in modulating red light signaling. Science 294, 1108–1111 (2001).
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).
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).
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).
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).
Song, C. et al. On the collective nature of phytochrome photoactivation. Biochemistry 50, 10987–10989 (2011).
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).
Gustavsson, E. et al. Modulation of structural heterogeneity controls phytochrome photoswitching. Biophys. J. 118, 415–421 (2020).
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).
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).
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).
Landgraf, F. T., Forreiter, C., Hurtado, P. A., Lamparter, T. & Hughes, J. Recombinant holophytochrome in Escherichia coli. FEBS Lett. 508, 459–462 (2001).
Hirose, Y. et al. Green/red cyanobacteriochromes regulate complementary chromatic acclimation via a protochromic photocycle. Proc. Natl Acad. Sci. USA 110, 4974–4979 (2013).
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).
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).
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).
Kabsch, W. XDS. Acta Crystallogr. D 66, 125–132 (2010).
Evans, P. R. & Murshudov, G. N. How good are my data and what is the resolution? Acta Crystallogr. D 69, 1204–1214 (2013).
Winn, M. D. et al. Overview of the CCP4 suite and current developments. Acta Crystallogr. D 67, 235–242 (2011).
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007).
Emsley, P. B., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D 66, 486–501 (2010).
Murshudov, G. N. et al. REFMAC5 for the refinement of macromolecular crystal structures. Acta Crystallogr. D 67, 355–367 (2011).
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).
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).
We thank SFB 1078 for funding (subprojects B6 to P.H. and B8 to J.H.). The X-ray diffraction measurements were carried out at BL14.1 at BESSY II (Helmholtz-Zentrum Berlin für Materialien und Energie (HZB)) and at ID30-A3 at ESRF with support from CALIPSOplus (Grant Agreement 730872 of the EU Framework Programme HORIZON 2020) and HZB. We thank L.-O. Essen (University of Marburg) for collaboration in the early stages of the project, W. Wende (Giessen) for cooperation in measuring the CD spectra and C. Lang (Giessen) for expert technical assistance. We dedicate this paper to the memory of Winslow Briggs.
The authors declare no competing interests.
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Left, Sorghum bicolor; Right Glycine max. Above: RR spectra of the Pr states (blue traces) and their photoconversion products (red traces) obtained upon 670 and 750 nm irradiation at ambient temperature. All spectra were measured at 90 K with 1064 nm excitation. The spectral regions labelled are indicative of (i) the methine bridge configurations and conformations (C=C stretching modes of the A-B and C-D methine bridges at ca. 1600–1650 cm−1), (ii) pyrrole nitrogen protonation state (N-H in-plane bending modes of rings B and C at ca. 1550 – 1580 cm−1) and (iii) the C-D methine bridge torsion (hydrogen-out-of-plane [HOOP] mode at ca. 795 - 825 cm−1). The broad feature at ca. 1460 cm−1 is largely due to non-resonant Raman bands of the protein. The high intensity of this feature relative to the RR bands of the chromophore indicates that the latter experience a low resonance enhancement. Below: IR “photoproduct minus Pr” difference spectra obtained upon irradiation with 670 and 750 nm at ambient temperature. The positive signals indicated by black lines and labels refer to the photoproduct, whereas the grey lines and labels mark the signals of the Pr state. Representative spectra based on at least two samples are shown. Spectra for each sample were measured several times. Each spectrum is based on 1000 separate FT scans.
Above: Superimposition of peptide chains with chromophores. The N- and C-termini, the two unresolved loops and the somewhat deviant R234-D236 regions are labelled. Inset: superimposition of the PCB (cyan) and P(B (green) D rings. Below: Chemical structures of PCB, PΦB, and the incorrect model used in 4OUR (PDB cofactor codes CYC, O6E and 2VO, respectively). Note that the uncharged structures are shown, whereas in both Pr and Pfr holoprotein states all four cofactor nitrogens are protonated.
Extended Data Fig. 3 Gm.phyA(PG)-PCB 2.1 Å structure (PDB 6TC7) of subunit B.
PCB (cyan), PAS (slate), GAF (gold), PCB (cyan), waters (red spheres). PW, pyrrole water. Subunit A (grey) is superimposed.
Extended Data Fig. 4 Gm.phyA(PG)-PCB 2.1 Å structure (PDB 6TC7) of subunit B including side chains and hydrogen bonding network.
Note that the weak (3.1 Å) D-ring carbonyl – H370 hydrogen bond in subunit B is somewhat stronger (3.0 Å) in subunit A. PCB (cyan), PAS (slate), GAF (gold), PCB (cyan), waters (red spheres). PW, pyrrole water.
Above. Superimposition of Gm.phyA(PG)-PCB subunit B (GAF domain, gold) with subunit A (grey) superimposed. Although the more mobile N-terminal section of the “380s” loop is missing (gold dashes), the final triad of the putative NLS (R360-R362) is resolved. Below. Alignment of the “380s” loop region in plant phytochromes (from Mathews et al. 47). sm, Selaginella martensii; cp, Ceratodon purpureus; ac, Adiantum caperis-veneris; atA-D, Arabidopsis thaliana PHYA-D; cpA, Curcurbita pepo PHYA; psA, Pisum sativum PHYA; stA/B, Solanum tuberosum PHYA/B, asA-D, Avena sativa PHYA; osA/B, Oryza sativa PHYA/B; zmA, Zea mais PHYA. The K(R/K)K(R/K) consensus is boxed red.
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Nagano, S., Guan, K., Shenkutie, S.M. et al. Structural insights into photoactivation and signalling in plant phytochromes. Nat. Plants 6, 581–588 (2020). https://doi.org/10.1038/s41477-020-0638-y
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