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. 2019 Nov 26;118(2):415–421. doi: 10.1016/j.bpj.2019.11.025

Modulation of Structural Heterogeneity Controls Phytochrome Photoswitching

Emil Gustavsson 1, Linnéa Isaksson 1, Cecilia Persson 2, Maxim Mayzel 2, Ulrika Brath 2, Lidija Vrhovac 1, Janne A Ihalainen 3, B Göran Karlsson 1,2, Vladislav Orekhov 1,2, Sebastian Westenhoff 1,∗
PMCID: PMC6976809  PMID: 31839260

Abstract

Phytochromes sense red/far-red light and control many biological processes in plants, fungi, and bacteria. Although the crystal structures of dark- and light-adapted states have been determined, the molecular mechanisms underlying photoactivation remain elusive. Here, we demonstrate that the conserved tongue region of the PHY domain of a 57-kDa photosensory module of Deinococcus radiodurans phytochrome changes from a structurally heterogeneous dark state to an ordered, light-activated state. The results were obtained in solution by utilizing a laser-triggered activation approach detected on the atomic level with high-resolution protein NMR spectroscopy. The data suggest that photosignaling of phytochromes relies on careful modulation of structural heterogeneity of the PHY tongue.

Significance

Phytochromes are light-sensing proteins found in plants, fungi, and bacteria. They are very primitive equivalents of human eyes and essential for an organism to change its growth or development in response to environmental light conditions. Limited experimental information about the dynamics and signaling mechanism for phytochromes exists. In this study, we investigate the structural dynamics using solution NMR spectroscopy. The pioneering experiments reveal that a specific element important for signaling, called the “PHY tongue,” transforms from a structurally heterogenous dark state, with two or more conformations present, to an ordered and homogenous light state. We suggest that, upon illumination, this transition locks the phytochrome in the activated state and that this explains how phytochromes modulate their photoactivity.

Introduction

Phytochromes are light-sensing proteins that monitor the level, intensity, duration, and color of environmental light (1, 2, 3). They control numerous light-dependent processes in plants, fungi, and bacteria (4, 5, 6, 7, 8). Phytochromes are promising targets in the expanding field of optogenetics to allow precise light control of various internal cellular processes (9). Absorption of red light by the dark-adapted (Pr) state leads to formation of the light-activated (Pfr) state. The dark state can be recovered by far-red light.

Most phytochromes are homodimeric proteins, with the common conserved domain architecture Per/Arndt/Sim-cGMP phosphodiesterase/adenyl cyclase/Fh1A-phytochrome specific (PAS-GAF-PHY) as the photosensory core module (10,11). A covalently linked bilin chromophore is attached via a thioether linkage to a conserved cysteine in either the PAS domain (bacteria) or GAF domain (cyanobacteria and plants) (12). In cyanobacteria and bacteria, the output domain often consists of a C-terminal histidine kinase, but in plants and fungi, a so-called N-terminal extension and C-terminal output domains, consisting of two PAS domains and a kinase-like domain, are present.

Phytochromes with a PAS-GAF-PHY photosensory module have a characteristic hairpin loop, called “the tongue” (residue 444–476), which extends from the PHY domain to the chromophore binding pocket in the GAF domain (Fig. 2). Crystallographic analysis has indicated that this tongue changes between two distinct structures from β-sheet configuration in dark to α-helical in light state (13, 14, 15). This refold has been connected to an opening of the dimer in crystallo and in solution, which most likely results in modification of the output domain. The tongue has, therefore, a key role in signal transduction (13).

Figure 2.

Figure 2

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Backbone chemical shift assignment of the dark (left) and light (right) state of the PAS-GAF-PHY photosensory module. Assigned residues (green) were mapped on structures with PDB: 4Q0J and 4O01, respectively. The biliverdin chromophore is colored in orange, and the helical spine is marked in the light state. The PHY tongue and its refolding between the dark and the light state is highlighted with a ring. To see this figure in color, go online.

The abovementioned structural models were built under the assumption that the protein is a static entity with well-defined states in dark and light. However, proteins are not static entities in living organisms; rather, their structures are highly dynamic. This is of functional relevance (16).

NMR spectroscopy is a powerful tool for examining the structure and dynamics of biological macromolecules, but NMR analysis of large proteins, like bacteriophytochromes, is difficult because of increased spectral complexity and sensitivity losses (17). These obstacles are probably the reason why solution-state NMR has only been applied to phytochrome PAS or PAS-GAF domains and not the complete photosensory core (18, 19, 20, 21, 22). The chromophore binding pocket has previously been studied using both solution- and solid-state NMR. With solid-state NMR, two hydrogen bond and charge distribution configurations have been detected in the chromophore-binding pocket of a cyanobacterial phytochrome in the dark-adapted state (23). Recent solution NMR of a chromophore-binding fragment of a red/green cyanobacteriochrome has confirmed this, in agreement with optical spectroscopy (18,24). Resonance Raman and time-resolved infrared spectroscopy have detected two chromophore conformations in the light state in cyanobacterial and plant phytochromes (25).

The evolutionarily conserved tongue region (residues 444–476) is in close contact with the chromophore, and the prevailing hypothesis is that upon illumination, distinct conformational changes in the chromophore-binding pocket are transduced to the tongue region, resulting in refolding (13,26, 27, 28). However, limited experimental information about the dynamic contribution of the tongue region of phytochromes exists. In this study, we aim to investigate the correlation of dynamics and photoactivation with a focus on the important tongue region in phytochromes. To do this, we used high-resolution protein solution NMR spectroscopy of the complete PAS-GAF-PHY photosensory module combined with a laser-triggered approach to control the photochemical state of the phytochrome.

Materials and Methods

The construct for the Deinococcus radiodurans monomeric phytochrome PAS-GAF-PHY fragment has previously been cloned (pET21b) and transformed into BL21(DE3) cells (29). The K177A, K476A, and K460A mutant constructs were made with the Quikchange Lightning Site-Directed Mutagenesis Kit (Agilent Technologies, Santa Clara, CA). The clones were verified by sequencing and transformed into competent BL21(DE3) cells.

Small-scale overnight cultures of the phytochrome were prepared in Luria-Bertani media. 4 mL of the cultures was inoculated to 200 mL of M9 media (Na2HPO4, KH2PO4, NaCl, NH4Cl, d-glucose, thiamine, MgSO4, CaCl2, H2O). 0.02 g of all unlabeled amino acids, except for lysine, were solubilized with M9 media and added to the cultures. Methionine and cysteine were solubilized with DTT and M9 media. To tryptophane and tyrosine, HCl and KOH were added, respectively, until the amino acids were completely solubilized. The cultures were grown at 37°C at 180 rpm. 15 min before induction, 0.2 g of all unlabeled amino acids, except for lysine, was added again, together with 20 mg of 13C,15N-labeled lysine. At OD 0.64, the temperature was decreased to 22°C, and 1 mM isopropyl β-D-1-thiogalactopyranoside was added. Induction continued for 24 h before the cells were harvested. An Emulsiflex C3 (Avestin, Ottawa, Canada) was used to disrupt the cells, followed by a 30-min centrifugation at 10,000 rpm. 3 mg of biliverdin (BV) hydrochloride (Frontier Scientific, Logan, UT) was dissolved in 1 mL of 30 mM Tris buffer (pH 8.0) with 8 M NaOH titrated until BV was completely solubilized. The cell lysates and the BV were incubated in the dark overnight at 4°C. The samples were filtered and loaded onto a 5-mL HisTrap HP column (GE Healthcare, Chicago, IL). The proteins were eluted with a linear gradient of imidazole with 30 mM Tris, 50 mM NaCl, and 300 mM imidazole (pH 8.0). The proteins were concentrated with a 30-kDa MWCO Vivaspin (Sartorius, Göttingen, Germany) to 3 mL and loaded to a HiLoad 16/600 Superdex 200-pg column (GE Healthcare, Chicago, IL) equilibrated with 30 mM Tris (pH 8.0). Eluted fractions were concentrated as described above. The buffer was exchanged by centrifugation to 25 mM NaPi and 50 mM NaCl (pH 6.9). UV-Vis was used to verify that the proteins could photoswitch, and the purity was checked using SimplyBlue Coomassie (Invitrogen, Waltham, MA) staining of Mini-PROTEAN TGX Stain-Free gel (Biorad, Hercules, CA). PageRuler Prestained Protein Ladder (Fermentas, Hanover, MD) was used as a standard.

Cells were adapted to D2O by streaking out on M9 agar plates containing increasing concentrations of D2O (25, 50, 75, and 100%). Colonies from the 100% D2O plate were picked, and 5 mL of overnight cultures of the phytochrome was prepared in M9 media (Na2HPO4, KH2PO4, NaCl, 15NH4Cl, u-13C-d-glucose, thiamine, MgSO4, CaCl2, D2O). 1 mL of one of these cultures was inoculated to 50 mL of M9 media. This culture was further used to inoculate 2 × 500 mL of M9 media. At OD 0.6, the temperature was decreased to 30°C, and 1 mM isopropyl β-D-1-thiogalactopyranoside was added. Induction continued for 22 h before the cells were harvested. The sample was purified according to the protocol stated above.

0.8 mM 13C,15N-lysine-labeled phytochrome samples (wild type, K476A, and K460A) was mixed with 10% D2O, 1× Complete EDTA-free (Roche, Basel, Switzerland), and 0.02% azide. 3-mm Shigemi tubes were used. To measure the phytochromes in the dark-adapted state, the samples in the NMR tube were illuminated in darkness using a 780-nm light-emitting diode (LED) (1.5 mW) for 10 min. The samples were injected into the magnet in darkness. To convert the samples to the light state, a 660-nm LED (14.5 mW) was used for illumination for 10 min.

0.8 mM of2H,13C,15N-labeled phytochrome sample was mixed with 10% D2O, 1× Complete EDTA-free (Roche), and 0.02% azide. A 3-mm Shigemi tube was used. To measure the phytochrome in the dark-adapted state, the sample in the NMR tube was illuminated in darkness using a 780-nm LED for 10 min. The sample was injected into the magnet in darkness. To convert the sample to the light state, a 660-nm LED laser (400 mW) was connected to an optical fiber with a diffuser tip (Laser components, Olching, Germany), which was inserted into the sample inside the 3-mm Shigemi tube (30). The sample was illuminated for 5 s every 20 min during data acquisition to continuously keep the sample in the light state.

NMR experiments were performed at 15, 25, 45, and 55°C on Bruker spectrometers with Larmor frequencies of 800 and 700 MHz equipped with 3-mm triple-resonance TCI cryoProbes. [1H,15N]-TROSYs were recorded for both the dark-adapted and light state for the wild type, the K460A mutant, and the K476A mutant. Visualization and verification of the assignment were performed using the CCPN software package (31).

NMR experiments were performed at 45°C on a Bruker spectrometer with a Larmor frequency of 800 MHz equipped with a 3-mm triple-resonance TCI cryoProbe. The automated backbone assignment was obtained using a previously reported assignment platform (32), which uses nonuniformly sampled three-dimensional (3D) TROSY-type experiments (HNCO, HNCA, HN(CO)CA, HNCACB, HN(CO)CACB, HN(CA)CO) combined with targeted acquisition (TA) and statistical validation. The details of this TA platform will not be discussed further in this article. [1H,15N]-TROSYs were recorded before and after all 3D spectra and also between each TA cycle to verify that the sample was identical throughout the entire measurement time. The 3D [15N]-NOESY-TROSYs were also recorded for both states and used during the manual assignment process. Verification of the automatically assigned residues and manual assignment were performed using the CCPN software package (31).

The secondary chemical shifts Δ δ = δ − δrc were obtained by subtracting the random coil value for each residue (δrc) from the chemical shift for the same residue (δ). The random coil data were obtained from Wishart and Sykes (33). The web server chemical shift index 3.0 was used to identify the location of secondary structure and random coil in the dark-adapted and light state of the phytochrome.

UV-Vis spectra were recorded at 45°C with scanning wavelengths of 280–850 nm. The sample was illuminated with a 780-nm LED for 10 min, and the spectrum was measured directly after (0 h). After a 72-h incubation in the dark, the UV-Vis spectrum was measured again.

Results

Backbone chemical shift assignment of the dark and light state

We report solution-state NMR data of the monomeric (509 residues) photosensory module (PAS-GAF-PHY) fragment of the DrBphP. The experiments were performed as described in Materials and Methods.

[1H15N]-TROSY spectra (Fig. 1) and a set of nonuniformly sampled 3D TROSY-type H-N-C spectra were recorded for both states. This set formed the basis for 1H,13C,15N backbone resonance assignment using TA (32,34), followed by manual inspection. In the dark spectra, 75% of all assignable residues were assigned for the dark-adapted state (Fig. 2). In the light spectra, we assigned 73% of all assignable residues (Fig. 2). Many assignments were transferred from the dark-adapted to the light-state spectra based on identical chemical shifts. Using our illumination conditions (660 nm), the light state is estimated to have an occupancy of 65% (13). Accordingly, our NMR spectra showed residual intensities of dark-adapted peaks in the “light” two-dimensional spectra (Fig. S1 A). However, in “light” 3D spectra, the intensities were below the detection limit for many of the dark-adapted signals. Therefore, the assignment of light-state signals was unambiguous for the majority of the peaks. In both states, unassigned residues cluster in the core region and the helical spine of the protein, likely because of poor proton back exchange. Attempts to favor exchange were performed by switching the protein between the dark and light states multiple times and at elevated temperatures, without any success.

Figure 1.

Figure 1

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The 2D [1H15N]-TROSYs for the dark and light state. (A) The 2D [1H15N]-TROSY of the dark-adapted state is shown. The assignments are annotated with one-letter amino acid codes and the sequence number. The crowded midregion is shown as an inset. BMRB: 27783. (B) Annotated 2D [1H15N]-TROSY of the light state is shown. The crowded midregion is shown as an inset. BMRB: 27784. To see this figure in color, go online.

Light-state contribution detected for the dark-adapted protein

Interestingly, the NMR data indicate that the dark-adapted proteins contain a contribution from the light state (Fig. S1 B). Peaks assigned for the light state were present in the “dark” spectra, and the intensities increased over time in darkness. Integration of multiple light-state peaks in the dark-detected [1H15N]-TROSY spectrum showed that the light-state peaks reached an intensity maximum of 10–15% of the equivalent dark-adapted-state peak. This is corroborated by UV-Vis spectra, which show an increase of far-red absorption when left in the dark, also corresponding to 10–15% light (Fig. S1, C and D). Our data thereby demonstrate directly that a light-state conformation exists even when DrBphP is illuminated with far-red light and left in the dark.

Structural heterogeneity in the PHY tongue region

Next, we turned our attention to the PHY tongue region. We could fully assign the tongue arms (residues 444–449 and 465–476) in the light state, including several highly conserved residues, but the tongue could not be resolved in the dark-adapted state (Fig. 2). We attribute the absence of tongue peaks in the dark-adapted state to peak broadening due to structural heterogeneity on a micro-to-millisecond timescale. This conclusion is rationalized by considering that the chemical shift of a nucleus can be altered when there is a conformational change in the protein. If the exchange between two conformations is slow, the different states of the nucleus will appear as two narrow peaks in the NMR spectrum. For a fast exchange, the peaks sharpen into a single peak at the averaged position. When the rate of interconversion is at an intermediate level, the peaks merge into one broad low-amplitude peak, beyond detection, at the population-averaged position. This occurs on the timescale of microseconds to milliseconds. The absence of tongue peaks in the dark state are thus an indication of structural heterogeneity with micro-to-millisecond exchange kinetics.

To strengthen this conclusion, we labeled all lysines in the protein with 13C and 15N (Fig. 3 A). K476 in DrBphPmon is located in the upper part of the tongue. One of the light-state peaks for the lysine-labeled sample (Fig. 3 B) disappeared upon mutation of K476 into alanine (circled), which identified the position of the K476 resonance in the spectrum (Fig. 3 C). The same lysine signal also disappeared when photoconverting the sample into the dark-adapted state. This strongly supports our interpretation that conformational heterogeneity exists in the PHY tongue in the dark-adapted state.

Figure 3.

Figure 3

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Lysine labeling of DrBphPmon. (A) The structure with lysines marked in red is shown (PDB: 4Q0J). (B) Shown is [1H15N]-TROSY of lysines, in which the tongue lysine disappears in the dark state because of structural dynamics on a micro-to-millisecond timescale. K460, which demonstrates two different conformations in light and one conformation in dark, is marked with arrows. (C) [1H15N]-TROSY of K476A mutant identifies the lysine located in the tongue, circled in red in both (B) and (C). To see this figure in color, go online.

To exclude rapid exchange with the bulk water as the cause for the loss of NMR signals in the PHY tongue in the dark-adapted state, we performed a number of test experiments. Factors that affect the rate of this exchange are hydrogen bonding, temperature, and pH. We therefore recorded NMR data for the lysine-labeled sample in the temperature range of 15–55°C, but we were not able to retrieve the lysine signal for the dark-adapted state (Fig. S2). Furthermore, K476 is also modeled as hydrogen bonded to the other β-strand in crystal structures, which should slow down the exchange with bulk water. Also, [1H15N]-TROSYs were recorded at pH 6.9, 6.0, and 4.8 because lowering the pH should slow down the rate of bulk-water exchange. None of the tested pH values could retrieve the K476 lysine signal in the tongue region (Fig. S3). This excludes the alternative explanation, strengthening our conclusion that structural heterogeneity of the PHY tongue is present.

The other solvent-exposed lysine (K460) was also assigned using a K460A mutant (Fig. S4). In the light state, this lysine appears as two separate peaks (arrows in Fig. 3 B). This is due to the presence of two separate states, which can be in slow exchange, whereas in the dark state, only one of the peaks appears. This indicates that K460 and K476 do not belong to the same structural element and that they have different structural dynamics. More experiments are needed to confirm this and to investigate the dynamic properties of the lysine (at 8.6 ppm) that show as a peak doubling in the dark state in the K476A mutant. A K177A mutant was also created, which precipitated most likely because of a destabilizing effect of the mutation.

Secondary chemical shift analysis of the PHY tongue in the light state

Variations in chemical shifts from random coil values (33), so-called secondary chemical shifts, provide insights into secondary-structure propensities. Positive values of Cα and CO and negative values of Cβ indicate α-helical structure, and the opposite indicates β-strand structure (35). A comparison of the secondary chemical shift calculated with TALOS-N using assigned chemical shifts (36) to the crystal structure (Fig. S5) shows an overall good agreement. In the light state (Fig. 4 A), the secondary chemical shifts for the tongue region strongly indicate not only an α-helical structure for the second arm (residue 465–476) but also the presence of a β-strand structure in the first arm (residue 444–449). Prediction of the secondary structures using the chemical shift index 3.0 web server (37) confirms this interpretation (see Fig. 4 B). Two stable secondary-structure elements for the tongue region in the light state are detected, which is in contrast to the conformationally heterogeneous dark state.

Figure 4.

Figure 4

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Secondary-structure elements in the light state for the two tongue arms in DrBphPmon. (A) Shown are the calculated secondary chemical shifts in light form, which correspond well with a β-strand structure in the first arm of the tongue, whereas the second arm is of an α-helical nature. Conserved residues are marked with an asterisk. (B) The predicted secondary-structure probability from H, NH, Cα, Cβ, and CO chemical shifts calculated by CSI 3.0 is shown (37). To see this figure in color, go online.

Discussion

For a long time, it was believed that the specific function of a protein is predetermined by its unique 3D structure. Proteins are, however, not solid rigid bodies, and the role of dynamics in the molecular mechanisms of these macromolecules has become increasingly clear.

We propose that the modulation of structural heterogeneity of the tongue region of DrBphPmon is of functional relevance (Fig. 5). Structural heterogeneity in phytochromes has been reported for residues surrounding the chromophore in smaller constructs, which excluded the PHY domain (18,23,24,38). The β-strand structure of the PHY tongue in the dark state, as detected by crystallography, should be considered as one of the conformations present in solution that is selected from two or more structures that preferentially crystallize. This conformational heterogeneity in the dark-adapted state can explain why DrBphP can adopt light-state-like crystals in darkness (39).

Figure 5.

Figure 5

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Transition from the structurally heterogeneous dark state to the ordered light state. Conformational fluctuation of the chromophore-binding pocket has previously been described for the dark state around the chromophore (18,23). Our data reveal that this structural heterogeneity extends far into the PHY tongue in DrBphP. Furthermore, we could show that the tongue adopts an ordered α-helix and β-strand structure in light state. We suggest that this transition locks the DrBphP in its photoactivated state. PDB: 4O0P (dark-adapted state) and 5C5K (light state). To see this figure in color, go online.

As such, the signaling mechanism does not simply involve a change of fold in the PHY tongue, as previously described (13,26, 27, 28), but instead, we suggest that the activity in DrBphP is triggered by altering the equilibrium between the structural heterogenous dark state toward the ordered state in the light. We suggest that the decreased conformational flexibility in the light state “locks” the phytochrome, altering the activity of the protein. This modulation of heterogeneity could be transferred to the helical spine and thereby further to the output domains (40).

We have performed a solution NMR analysis of the large system of DrBphPmon, gaining insight into the important signaling tongue region using a laser-triggered approach, exciting the protein inside the NMR magnet in a noncontinuous way during acquisition. Our study demonstrates that it is critical to study phytochromes in solution to reveal both the structure and the dynamic behavior of the protein in different functional states. Our first partial assignment of a complete photosensory module of DrBphPmon, enables further studies of the protein with atomic precision in solution, opening up for resolving the structural and dynamical mechanism of photosignaling in phytochromes.

Conclusion

We conclude that the PHY tongue region undergoes a transition from a structural heterogenous state to an ordered state upon light activation. Because the dynamics of the PHY tongue are directly controlled by the light conditions, we suggest that the alteration of the structure equilibrium toward the ordered light state is of biological significance for the photomodulation of phytochromes.

Author Contributions

E.G. and L.I. planned the project together with S.W., C.P., V.O., B.G.K., U.B., and J.A.I. E.G., L.I., and L.V. produced and purified all samples and performed all measurements and analyses together with V.O., M.M., and C.P. E.G., C.P., and L.I. performed the backbone assignment. E.G., L.I., and S.W. wrote the manuscript, with edits by J.A.I.

Acknowledgments

We thank Sami Mäkelä, Marjo Haapakoski, and Ilkka Minkkinen for initial temperature stability experiments performed with UV-Vis spectrophotometry. We also thank Michal Maj and Magnus Wolf-Watz for feedback for the manuscript.

S.W. thanks the Knut and Alice Wallenberg Foundation for an Academy Fellowship. J.A.I. acknowledges the Academy of Finland grant 296135 and Jane and Aatos Erkko Foundation. V.O. thanks the support by the Swedish Research Council (Research Grant 201504614). NMR spectroscopy was carried out at the Swedish NMR Centre at University of Gothenburg, supported by the Knut and Alice Wallenberg Foundation (NMR for Life) and the Science for Life Laboratory (SciLifeLab).

Editor: Monika Fuxreiter.

Footnotes

Emil Gustavsson and Linnéa Isaksson contributed equally to this work.

Supporting Material can be found online at https://doi.org/10.1016/j.bpj.2019.11.025.

Supporting Material

Document S1. Figs. S1–S5
mmc1.pdf (359.9KB, pdf)
Document S2. Article plus Supporting Material
mmc2.pdf (3.3MB, pdf)

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Figs. S1–S5
mmc1.pdf (359.9KB, pdf)
Document S2. Article plus Supporting Material
mmc2.pdf (3.3MB, pdf)

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