Plants adapt to the availability of light throughout their lives because it regulates so many aspects of their growth and reproduction. To detect the level of light, plant cells use proteins called phytochromes, which are also found in some bacteria and fungi. Phytochrome proteins change shape when they are exposed to red light, and this change alters the behaviour of the cell. The red light is absorbed by a molecule known as chromophore, which is connected to a region of the phytochrome called the PHY-tongue. This region undergoes one of the key structural changes that occur when the phytochrome protein absorbs light, turning from a flat sheet into a helix.

Claesson, Wahlgren, Takala et al. studied the structure of a bacterial phytochrome protein almost immediately after shining a very brief flash of red light using a laser. The experiments revealed that the structure of the protein begins to change within a trillionth of a second: specifically, the chromophore twists, which disrupts its attachment to the protein, freeing the protein to change shape. Claesson, Wahlgren, Takala et al. note that this structure is likely a very short-lived intermediate state, which however triggers more changes in the overall shape change of the protein.

One feature of the rearrangement is the disappearance of a particular water molecule. This molecule can be found at the core of many different phytochrome structures and interacts with several parts of the chromophore and the phytochrome protein. It is unclear why the water molecule is lost, but given how quickly this happens after the red light is applied it is likely that this disappearance is an integral part of the reshaping process.

Together these events disrupt the interactions between the chromophore and the PHY-tongue, enabling the PHY-tongue to change shape and alter the structure of the phytochrome protein. Understanding and controlling this process could allow scientists to alter growth patterns in plants, such as crops or weeds.

Phytochrome photosensor proteins are crucial for the optimal development of all vegetation on Earth (Butler et al., 1959; Gan et al., 2014; Quail et al., 1995). Prototypical phytochromes can exist in two photochemical states with differential cellular signaling activity, called red light-absorbing (Pr) and far-red light-absorbing (Pfr) state (Figure 1—figure supplement 1). As a result, phytochromes can distinguish two colors of light, providing plants, fungi, and bacteria with primitive two-color vision. Light is detected by a bilin chromophore, which is covalently linked to the photosensory core of the protein (Wagner et al., 2005), comprising of PAS (Per/Arndt/Sim), GAF (cGMP phosphodiesterase/adenyl cyclase/FhlA) and PHY (phytochrome-specific) domains. Two propionate side chains additionally anchor the chromophore non-covalently to the protein (Figure 1b). The signaling sites of the phytochrome are found in its C- and N-terminal output domains, which vary between species. Important for the signaling is a stretch of amino acids in the PHY domain, called the PHY-tongue, which changes from a β-sheet in Pr into an α-helix in Pfr state (Essen et al., 2008; Yang et al., 2008; Takala et al., 2014; Sanchez et al., 2019) The chromophore connects to the PHY-tongue via a strictly conserved aspartatic acid, which is expected to play a crucial role in signal transduction.

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Key to phytochrome function is the primary photoresponse on picosecond time scales. Here, light signals are translated into conformational changes. The changes arise in the electronically excited bilin, but must then be transduced to the surrounding protein residues. This prepares the protein for a formation of the first intermediate (Lumi-R for prototypical phytochromes), in which isomerization of the D-ring has likely occurred (Rüdiger et al., 1983; Dasgupta et al., 2009; Yang et al., 2012; Rockwell et al., 2009; Ihalainen et al., 2018). The mechanism that leads to the first intermediate is currently not well understood, because crystallographic observations of phytochromes directly after photoexcitation have not been available.

To address this gap of knowledge, we recorded time-resolved serial femtosecond X-ray crystallographic (SFX) data of the PAS-GAF domains of the phytochrome from Deinococcus radiodurans (DrBphP_CBD) at 1 ps and 10 ps after femtosecond optical excitation. The experiments were performed in Japan, using the SPring-8 Angstrom Compact Free Electron Laser (SACLA) tuned to 7 KeV (Tono et al., 2015). For homogeneous excitation of the crystals, we photoexcited micrometer-sized crystals in a grease jet with a photon density of 1.7 mJ/mm^₂ ($1/e^2$ measure, see Materials and methods) into the flank of the absorption peak at 640 nm (Figure 1—figure supplement 1). Taking into account the significant light scattering in the grease-buffer mixture (Figure 1—figure supplement 1), we estimate that the average number of photons per chromophore is 0.5–1 (see Materials and methods). We recorded the SFX data at 1 ps for several excitation fluences (Figure 2). Lowering the excitation density tenfold from 1.7 mJ/mm^₂ photons to 0.2 mJ/mm^₂ resulted in a joint reduction of all difference signals. Critical signals, like the twist of the D-ring and the photodissociation of the pyrrole water from the chromophore sustained when lowering the excitation densities, indicating that the signal arises predominately from one-photon excitation. The refined structure in dark (DrBphP_dark), 2.07 Å resolution (Table 1), was very similar to our previous dark structure solved by SFX (5K5B, RMSD 0.646 Å and 0.610 Å for monomers A and B) (Edlund et al., 2016), but the present crystals contained two monomers in the asymmetric unit (Figure 1a, Figure 1—figure supplement 2, Figure 1—figure supplement 3). The refined 1 ps structure was solved to 2.21 Å (Table 1).

Figure 2

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From the time-resolved data, we calculated Fourier difference electron density maps (${|F_o|}^{light}-{|F_o|}^{dark}$), which report on the change of structure due to optical excitation (see Materials and methods). Briefly, the diffraction data for light and dark were scaled to each other and subtracted, assuming preservation of the phases (see Materials and methods for details). The map at 1 ps indicates many significant changes in difference electron density (Figure 1a) above the background level of 3.0 standard deviations (σ) (Figure 1—figure supplement 4). The changes cluster around the chromophore, with the strongest negative densities for the pyrrole water (monomer A: −8.2σ, B: −9.4σ, Table 2). The map at 10 ps contains similar significant features, but at weaker overall intensity (pyrrole water A: −5.0σ, B:−6.5σ) (Figure 1—figure supplement 4 and Figure 1—figure supplement 5). We ascribe this to a lower population of the activated state at 10 ps compared to 1 ps. Monomer A has a lower signal strength than monomer B, but provided a clearer difference map around the chromophore. We refined a structural model (DrBphP_1ps) using extrapolated structure factors (Figure 3—figure supplement 1; Pande et al., 2016). The refinement of the structure against the 1 ps data was successful using a photoexcitation density of 8%. However, we aborted our attempts to refine a structural model against the 10 ps data, as the model would have become unreliable due to an even lower photoactivation yield. We focus our discussion on monomer A and the 1 ps time point, although all conclusions are supported by monomer B and the features observed in the difference maps at 10 ps (Table 2, Figure 1—figure supplement 5).

First, we inspect the D-ring region at 1 ps (Figure 3). We observe strong negative difference density features on the atoms of the D-ring (marked I, II, III), correlating with density gains at both faces of the ring (IV, V, VI). These features strongly indicate that the D-ring twists. The positive feature IV homes the N-H and C = O groups, whereas V and VI indicate densities for the methyl and vinyl groups in the twisted ring (Figure 3c). Excellent agreement was obtained between the observed difference map and the difference map calculated from DrBphP_1ps ($F_c^{1ps}-F_c^{dark}$), when the D-ring twists (C14-C15-C16-ND) from around 20° in the dark to 60° monomer A) and 90° monomer B) at 1 ps (Figure 3c and e). Although the twisting movement is clearly indicated by the difference map, we judge the precision of the angle to be low and approximately ±25°.

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Concomitant with the twist of the D-ring, the C-ring translates by approximately 0.69 Å as indicated by the correlated negative (VII) and positive (VIII) electron density (Figure 3a). Furthermore, the C-ring propionate chain detaches from its conserved anchoring residues Ser272 and Ser274 (IX and X, Figure 3a). The strictly conserved His260 retracts from its position (XI and XII) and Tyr263 moves upward at 1 ps (XIII and XIV, Figure 3b). The water network connecting the C-ring propionate, the D-ring C = O, and His290 rearranges accordingly (Figure 3a). The excellent agreement between calculated and observed difference maps confirms these observations (Figure 3d and e, Figure 3—figure supplement 2). We conclude that the twist of the D-ring causes detachment of the C-ring propionate from the protein scaffold by dislocation of the C-ring, facilitated by the associated hydrogen bonding network.

Turning our attention to the B-ring, we find that the B-ring propionate breaks its salt bridge to Arg254 (Figure 3—figure supplement 3). However, this is not caused by movements of the chromophore backbone, as we observe little change on the B-ring itself. Instead, we find that a water bridge between the B- and C-ring propionates is broken as indicated by negative difference electron densities on the waters (Figure 3—figure supplement 3). Additionally, the highly conserved helix from Ser257 to Val269, moves away from the chromophore by an average of 0.36 Å in monomer A and 0.62 Å in monomer B (distances relative to the pyrrole water, Figure 3—figure supplement 4). The changes of the D-ring are transduced to Ser257 via the side chains of His260 and Tyr263, and as a result, the hydrogen bond of Ser257 to the B-ring propionate group breaks. The amino acids in the stretch are over 50% conserved (Figure 3—figure supplement 4), suggesting that it has evolved to transfer an ultrafast signal. We conclude that relaxation of the protein is necessary for the detachment of the B-ring propionate from the protein scaffold.

Next to the changes around the D-ring, the maps reveal strong difference electron density on the A-ring (XVIII and XIX), Asp207 (XX to XXIII) (Figure 4a) and the pyrrole water (XV) (Figure 4b). When interpreted and modelled as downward movement of the A-ring and Asp207 and photodissociation of the pyrrole water from the chromophore, excellent agreement between calculated and observed difference electron density is obtained (Figure 4—figure supplement 1). The A-ring is covalently attached to the protein backbone in phytochromes (Song et al., 2014), which renders complete isomerization impossible, but is sufficiently flexible to accommodate the proposed changes. The pyrrole water may either move to feature (XVI), or occupy an anisotropic, worm-shaped feature which extends from the A-ring to the D-ring (XVII) (Figure 4b).

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Furthermore, correlated negative and positive electron density features are observed on backbone atoms of the highly conserved stretch from Pro201 to His209. These difference electron density features indicate that the residues move away from the centre of the chromophore by an average of 0.54 Å and 0.57 Å in monomers A and B, respectively (Figure 3—figure supplement 4b). The stretch includes Asp207 and it is located between the A-ring and the PHY domain, which makes it plausible that the changes in the chromophore cause this protein rearrangement. The changes are complemented by significant rearrangements of a stretch of waters and a conserved Tyr176 (Figure 4c).

The structure of DrBphP_CBD 1 ps after photoexcition reveals changes of the biliverdin chromophore and the surrounding residues. We find a twist of the D-ring, displacement of the C-ring, and associated changes of the water network which connects the D-ring, the C-ring propionate, and His290. Further, we identify a disruption of the salt bridge between the B-ring propionate and the Arg254, and significant changes around the A-ring, Asp207 and the pyrrole water. The changes are retained at 10 ps, even though at a lower population (Figure 1—figure supplement 5). The extensive and coordinated structural changes in the binding pocket (Figure 3) manifest a liberation of the chromophore from the protein scaffold, which we propose to be necessary for the conformational rearrangements to occur in the downstream photoconversion to Pfr.

Infrared spectral data indicate significant reorganization of the chromophore and several amino acids including the PHY-tongue region as early as in Lumi-R state, which is the first known ground state intermediate in the photoconversion from Pr to Pfr (Ihalainen et al., 2018; van Thor et al., 2007). However, the structure of the bilin and the binding pocket in Lumi-R is not known, because structural information is missing. Since the quantum yield of reaching the Lumi-R state is low (on the order of 10%), spectroscopic investigation of the mechanism is difficult, and it is currently not fully established how the Lumi-R state is reached. Crystallographic data does not report on whether the chromophore is electronically excited or not, hence we cannot determine whether the structure that we observe is in a relaxed excited state or in a ground state. The time delay (1 ps) supports that our structure presents an intermediate enroute to the Lumi-R state.

The D-ring of the bilin chromophore isomerizes around the C15-C16 bond from (Z) in Pr to (E) in Pfr (Rüdiger et al., 1983; Burgie et al., 2016; Takala et al., 2014; Yang et al., 2008; Essen et al., 2008). Circular dichroism spectroscopy and solid-state NMR spectroscopy have indicated that the position of the D-ring inverts from an ’$\alpha$’-facial (Pr) to a ’$\beta$’-facial (Pfr) position in cyanobacterial and plant phytochromes, whereas it stays ’$\alpha$’-facial in bacterial phytochromes (Rockwell et al., 2009; Song et al., 2011; Song et al., 2018). Based on anticipated steric clashes with the C-ring methyl group, it has been proposed that the D-ring rotates counter-clockwise in plant and cyanobacterial phytochromes, but clockwise in bacterial phytochromes (Rockwell et al., 2009). Moreover, spectroscopy has shown that the D-ring of the bilin chromophore is already isomerized in the Lumi-R state (van Thor et al., 2007; Heyne et al., 2002; Yang et al., 2012).

Seemingly contradictory, we now observe that the D-ring is rotated counter-clockwise by tens of degrees for the bacterial DrBphP at 1 ps time delay (Figure 3c and e). The conformation is strongly supported by the difference map. It contains two positive peaks (V and VI in Figure 3c), which indicate the new position of the vinyl and methyl group of the D-ring. We tested models in which the D-ring was rotated in a clockwise direction, but the agreement with the experimental difference map decreased. Thus, it may be that the D-ring indeed rotates counter-clockwise in bacterial phytochromes, similar to plant and cyanobacterial phytochromes. For complete isomerization, this would mean that the C-ring moves out of the way during the rotation. We observe significant movements of the C-ring, which may be an indication for that such a mechanism is possible. Raising a note of caution, we cannot fully exclude that the truncation of our phytochrome construct or the crystal packing influences the direction of rotation. NMR studies have reported conformational heterogeneity in the chromophore binding pocket of phytochromes in solution (Song et al., 2011; Lim et al., 2018; Song et al., 2018; Gustavsson et al., 2020). Crystallization could select one of the conformations, which may have a preferred rotation in the counter-clockwise direction. More experiments are needed to clarify this question.

It is interesting to compare the structural changes at 1 ps time delay to the changes observed in the conversion between Pr and Pfr (Burgie et al., 2016; Takala et al., 2014; Stojković et al., 2014). Major changes include a flipped D-ring, changes in conserved residues of the chromophore-binding pocket, for example Tyr176, His201 and Phe203, and refolding of the PHY tongue. The PHY tongue is not included in our construct, but Tyr176 and Phe203 are associated with difference electron density features in our maps (Table 2). However, the movements are much smaller at 1 ps compared to the Pr-to-Pfr transition. This is not unexpected, given the short time delays, but it shows that the residues are tightly coupled to the chromophore. Interestingly, the Pr and Pfr structures also reveal a sliding movement of the entire chromophore (Yang et al., 2011; Burgie et al., 2016; Takala et al., 2014). This requires that the propionic groups have to break their bonds to the protein scaffold. Our data indicate that this is part of the primary photoresponse.

The photodissociation of the pyrrole water from the chromophore is a surprising finding. The pyrrole water is ubiquitously found in phytochrome structures (Essen et al., 2008; Yang et al., 2008; Otero et al., 2016; Burgie et al., 2016; Wagner et al., 2005; Burgie et al., 2014; Schmidt et al., 2018; Yang et al., 2011). Our fluence dependent SFX data show that the negative density on the pyrrole water is the last signal to disappear when lowering the photon excitation densities 10-fold (Figure 2). This makes us confident that the photodissociation reaction is not caused by multi-photon effects. The removal of the water requires significant energy, because the hydrogen bonds to the A-, B-, and C-rings of the chromophore and the backbone C = O group of Asp207 have to be broken. We do not think that the twist of the D-ring causes this through direct steric interactions, because there is no contact between the pyrrole water and the D-ring. Rather, it may be triggered by an excited state charge redistribution between the pyrrole water and the chromophore, for example by ultrafast proton or electron transfer (Toh et al., 2010). Such charge re-distributions are typically facilitated by changes in geometry (Nosenko et al., 2008) and may therefore be caused indirectly by structural changes of the A-, C-, or D-rings, but this requires further investigation.

Conformational changes of the A-ring, Asp207 and the pyrrole water have not been considered to occur on picosecond time scales. The strictly conserved Asp207 is a key residue for signal transduction because it connects the chromophore to the PHY-tongue in Pr and Pfr (Essen et al., 2008; Yang et al., 2008; Takala et al., 2014). Its displacement suggests, together with the relocation of the residue stretch surrounding it, that disruption of the GAF-PHY interface may occur as early as 1 ps after photoexcitation (Figure 3—figure supplement 4b). With a hydrogen bond to the pyrrole water and in tight steric contact with the A-ring, Asp207 thereby acts as an extended arm of the chromophore. We propose that the photodissociation of the pyrrole water from the bilin and the change of the A-ring are integral parts of ultrafast phytochrome signaling toward the PHY domain.

We demonstrate that within 1 ps, the D-ring twists, that the chromophore is liberated from the protein (Figure 5a) and that movements of the pyrrole water, the A-ring and Asp207 lead to signaling directed toward the PHY-tongue (Figure 5b). When mapped on the structure of the complete photosensory core module (Takala et al., 2014), both changes work together to destabilize the Arg466:Asp207 salt bridge. Tyr263 moves up, caused by the twist of the D-ring, and Asp207 moves down, caused by changes of the A-ring, retracting both residues from the salt bridge.

Figure 5

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Our data reveal a highly collective primary photoresponse for phytochromes. This is consistent with the fact that most point mutations of conserved residues alter, but do not inhibit, photoconversion (Wagner et al., 2008). The ultrafast structural changes are more extensive than in bacteriorhodopsin, photoactive yellow proteins, and in a fluorescent protein (Pande et al., 2016; Nogly et al., 2018; Coquelle et al., 2018). While previously observed ultrafast backbone movements have been interpreted as ’protein quakes’ for myoglobin and bacteriorhodopsin (Barends et al., 2015; Nogly et al., 2018; Toh et al., 2010), the present backbone motion in the phytochrome binding pocket are much more directed (Figure 3—figure supplement 4). The changes occur in highly conserved regions of the protein and are part of the collective signaling response of the entire binding pocket.

Phytochromes have to be able to stabilize the bilin and to direct its photoisomerization from two photochemical ground states, Pr and Pfr. These differ both structurally and electronically, which precludes a single reaction trajectory for isomerization in the two directions. With this in mind, the observed primary photoresponse is reasonable. The structural signal is highly delocalized already at 1 ps, causing near-simultaneous liberation of the chromophore and initial signal transduction. We propose that these reaction trajectories stabilize each other, navigating the protein into a productive reaction path. The multidimensional reaction trajectory is consistent with the low quantum yields for photoconversion (Lamparter et al., 1997), which are characteristic for the phytochrome superfamily. Whereas the twisting motion of the D-ring has been the working model for phytochrome activation and is now confirmed, the photodissociation of the pyrrole water is highly surprising. We propose that both chemical events work together and enable phytochrome proteins to translate light information into structural signals, guiding the growth and development of plants, fungi, and bacteria on Earth.

Crystallography data have been submitted to protein data bank (PDB) dark:ID: D_1292104678 and PDB ID: 6T3L 1ps:ID: D_1292104679 and PDB ID: 6T3U. Raw diffraction images are deposited under ID 121 at CXIDB.

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Author details

1. Elin Claesson

   Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

   Contribution

   Planned experiments, Purified the protein and prepared the microcrystals, Acquired the data at SACLA, Computed difference electron density maps and refined the structural models, Wrote the paper with input from all authors

   Contributed equally with

   Weixiao Yuan Wahlgren and Heikki Takala

   Competing interests

   No competing interests declared

2. Weixiao Yuan Wahlgren

   Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

   Contribution

   Purified the protein and prepared the microcrystals, Acquired the data at SACLA, Computed difference electron density maps and refined the structural models

   Contributed equally with

   Elin Claesson and Heikki Takala

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0413-165X

3. Heikki Takala

   1. Department of Biological and Environmental Science, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland
   2. Department of Anatomy, Faculty of Medicine, University of Helsinki, Helsinki, Finland

   Contribution

   Purified the protein and prepared the microcrystals, Acquired the data at SACLA, Computed difference electron density maps and refined the structural models

   Contributed equally with

   Elin Claesson and Weixiao Yuan Wahlgren

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2518-8583

4. Suraj Pandey

   Physics Department, University of Wisconsin-Milwaukee, Milwaukee, United States

   Contribution

   Acquired the data at SACLA and processed the crystallography data

   Competing interests

   No competing interests declared

5. Leticia Castillon

   Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

   Contribution

   Purified the protein and prepared the microcrystals, Acquired the data at SACLA

   Competing interests

   No competing interests declared

6. Valentyna Kuznetsova

   Department of Biological and Environmental Science, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

   Contribution

   Acquired the data at SACLA, Measured the UV-Vis spectroscopic data

   Competing interests

   No competing interests declared

7. Léocadie Henry

   Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

   Contribution

   Acquired the data at SACLA

   Competing interests

   No competing interests declared

8. Matthijs Panman

   Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

   Contribution

   Acquired the data at SACLA

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3853-123X

9. Melissa Carrillo

   Department of Biology, Northeastern Illinois University, Chicago, United States

   Contribution

   Acquired the data at SACLA

   Competing interests

   No competing interests declared

10. Joachim Kübel

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

11. Rahul Nanekar

    Department of Biological and Environmental Science, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

12. Linnéa Isaksson

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

13. Amke Nimmrich

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

14. Andrea Cellini

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

15. Dmitry Morozov

    Department of Chemistry, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

    Contribution

    Provided the geometry restrains biliverdin in the excited state

    Competing interests

    No competing interests declared

16. Michał Maj

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

17. Moona Kurttila

    Department of Biological and Environmental Science, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

    Contribution

    Measured the UV-Vis spectroscopic data

    Competing interests

    No competing interests declared

18. Robert Bosman

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

19. Eriko Nango

    1. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    2. RIKEN SPring-8 Center, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

20. Rie Tanaka

    1. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    2. RIKEN SPring-8 Center, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

21. Tomoyuki Tanaka

    1. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    2. RIKEN SPring-8 Center, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

22. Luo Fangjia

    1. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    2. RIKEN SPring-8 Center, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

23. So Iwata

    1. Department of Cell Biology, Graduate School of Medicine, Kyoto University, Kyoto, Japan
    2. RIKEN SPring-8 Center, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

24. Shigeki Owada

    1. RIKEN SPring-8 Center, Hyogo, Japan
    2. Japan Synchrotron Radiation Research Institute, Hyogo, Japan

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

25. Keith Moffat

    Department of Biochemistry and Molecular Biology and Institute for Biophysical Dynamics, University of Chicago, Chicago, United States

    Contribution

    Conceived the experiments

    Competing interests

    No competing interests declared

26. Gerrit Groenhof

    Department of Chemistry, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

    Contribution

    Provided the geometry restrains biliverdin in the excited state

    Competing interests

    No competing interests declared

27. Emina A Stojković

    Department of Biology, Northeastern Illinois University, Chicago, United States

    Contribution

    Acquired the data at SACLA

    Competing interests

    No competing interests declared

28. Janne A Ihalainen

    Department of Biological and Environmental Science, Nanoscience Center, University of Jyvaskyla, Jyvaskyla, Finland

    Contribution

    Conceived the experiments, Acquired the data at SACLA

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-8741-1587

29. Marius Schmidt

    Physics Department, University of Wisconsin-Milwaukee, Milwaukee, United States

    Contribution

    Conceived the experiments, Acquired the data at SACLA, Computed difference electron density maps and refined the structural models, Wrote the paper with input from all authors

    For correspondence

    smarius@uwm.edu

    Competing interests

    No competing interests declared

30. Sebastian Westenhoff

    Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden

    Contribution

    Conceived the experiments, Planned experiments, Acquired the data at SACLA, Computed difference electron density maps and refined the structural models, Wrote the paper with input from all authors

    For correspondence

    sebastian.westenhoff.2@gu.se

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-6961-8015

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