Photoreaction Dynamics of LOV1 and LOV2 of Phototropin from

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Photoreaction Dynamics of LOV1 and LOV2 of Phototropin From Chlamydomonas Reinhardtii Yusuke Nakasone, Masumi Ohshima, Koji Okajima, Satoru Tokutomi, and Masahide Terazima J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.7b10266 • Publication Date (Web): 22 Jan 2018 Downloaded from http://pubs.acs.org on January 22, 2018

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The Journal of Physical Chemistry

Photoreaction of Cr-Phototropin constructs

Photoreaction Dynamics of LOV1 and LOV2 of Phototropin from Chlamydomonas Reinhardtii Yusuke Nakasone,† Masumi Ohshima,† Koji Okajima,# Satoru Tokutomi,§ and Masahide Terazima*,† †

Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan # Graduate School of Science and Technology, Keio University, Kanagawa 223-8522, Japan § Department of Biological Science, Graduate School of Science, Osaka Prefecture University, Sakai, Osaka 599-8531, Japan Running title: Photoreaction of Cr-Phototropin constructs

* To whom correspondence should be addressed: Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan Tel:81-75-753-4026, FAX: 81-75-753-4026, e-mail: [email protected]

ABSTRACT Phototropin is a blue light sensor protein found in higher plants and green algae. Photochemical reactions of a variety of differently truncated constructs of a phototropin from Chlamydomonas reinhardtii (Cr) (LOV1, LOV1-hinge, LOV2, LOV2-linker and hinge-LOV2) are investigated. In the dark state, LOV1 is in dynamic equilibrium between the monomer and dimer, and the main photochemical reaction is dimerization of the monomer and dissociation of the dimer. On the other hand, LOV1-hinge exists as the monomer and the photochemical reaction is the dimerization reaction associated with the unfolding of the helix of the hinge domain. LOV2 in the dark state is monomeric. The conformation changes after the photoexcitation of LOV2, and LOV2-linker are minor, which differs noticeably from the reaction of LOV2-Jα and LOV2-linker from Arabidopsis thaliana (At). The linker region including the Jα helix is rather stable upon photoexcitation. The helix of the hinge domain of hinge-LOV2 is slightly unfolded in the dark state and the major photoreaction is the dimerization event. The dark recovery rate of LOV2 was found to decrease significantly in the presence of the hinge domain. These photochemical properties of Cr phot are considerably different from those of At phot regarding conformational changes and their kinetics, although Cr phot has been reported to rescue the phot function in At. The differences and the diversity of phots are discussed.

INTRODUCTION Phototropin (phot) is a blue light sensor protein found in higher plants and green algae 1. Higher 1 ACS Paragon Plus Environment

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Photoreaction of Cr-Phototropin constructs plants often possess two isoforms of phot (phot1 and phot2), and these two isoforms regulate phototropism, stomatal opening, leaf flattening and chloroplast movement to optimize photosynthesis

2–6

. The unicellular

green alga Chlamydomonas reinhardtii (Cr) has one phot, which triggers blue-light-dependent sexual differentiation, photosynthetic gene expression and photoprotection, and regulates eyespot size and phototaxis 7–11. Phots consist of two light-oxygen-voltage domains (LOV1 and LOV2) and a serine/threonine kinase domain 2,7. These domains are connected by hinge and linker domains as shown in Fig. 1(a). The two LOV domains are light-sensing modules, and the activity of the kinase domain is regulated by the LOV domains in a light-dependent manner to control downstream outputs and signaling events. LOV has a three-dimensional structure composed of several α-helices and β-strands

12–14

. In the

dark, LOV binds a flavin mononucleotide (FMN) as a chromophore non-covalently in a hydrophobic binding pocket formed by the α/β-scaffold (Fig. 1(b)). Upon photoexcitation, LOV undergoes a photoreaction cycle 15–17

through the transient formation of a cysteinyl adduct between the FMN and a nearby cysteine residue

.

The adduct form, which is characterized by an absorption spectral change (Fig. 1(c) and (d)), thermally reverts to the initial state in a time range of seconds to minutes 18. The photoreaction in the vicinity of the chromophore (the absorption spectral change) is well conserved among all LOV domains studied and adduct formation is a triggering step leading to conformational changes in the protein that stimulate signal transduction. Photochemical and photophysical properties of phots from higher plants such as Arabidopsis thaliana (At) have been studied widely. Despite similarities in their structures and photoreaction cycles, LOV1 and LOV2 have distinct roles in regulating phot activities. In At-phot, LOV1 attenuates the light sensitivity of phot, whereas LOV2 is primarily responsible for the light-dependent phosphorylation of the kinase domain and is indispensable for signal transduction mediated by At-phot 19–21. Since LOV2 is vital for the function of phot, the reaction dynamics of constructs containing the LOV2 domain have been studied extensively by several biophysical techniques, including nuclear magnetic resonance (NMR), Fourier transform infrared spectroscopy (FTIR), circular dichroism (CD), small angle X-ray scattering (SAXS) and transient grating (TG)

22–29

. A helix, located in the C-terminal region of the

LOV2 domain (Jα-helix: a part of the linker domain), was found to contribute to light induced kinase activation. The Jα-helix embeds on the surface of the LOV2 core and is located on the opposite side of the FMN binding pocket in the dark, and unfolds in the light state

30

. Time-resolved detection of the

conformational changes of several constructs of phot from At have been reported by the TG method, which can detect protein reaction dynamics of secondary/tertiary structural changes and changes in the oligomeric state (association/dissociation) from the view point of a change in the diffusion coefficient (D)

31–33

. For

example, photoexcitation of the At phot1LOV2-linker domain results in the dissociation of the linker from the LOV2 core with a time constant of 300 µs and the subsequent unfolding of the linker helix (1 ms) 27. The dissociation and subsequent unfolding are important steps for kinase activation and are conserved in At phot2LOV2-linker with slight differences in the time constant (140 µs and 2 ms for dissociation and unfolding, respectively) 26. Moreover, the A'α helix located on the N-terminal side of the LOV2 domain was shown to unfold with a time constant of 12 ms 28. The signal propagation in phot presumably is achieved by the unfolding of these flanking helices of LOV2. 2

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Photoreaction of Cr-Phototropin constructs The LOV1 domain of At phots is postulated to act as a dimerization site, because dimeric forms have been observed by crystallography for At phot1LOV1 and At phot2LOV1

34

. Size exclusion

chromatography (SEC) measurements also confirmed that At phot1LOV1 exists as a stable dimer and At phot2LOV1 was shown to be in dynamic equilibrium between the monomer and dimer states in the dark 35,36. Using the TG method, the dimer of At phot1LOV1 was shown to associate to form a tetramer and the monomer of At phot2LOV1 dimerizes upon blue light illumination

35,36

. When the C-terminal region of At

phot2LOV1 is included (At phot2LOV1-hinge), no secondary structural change is observed 35. These results suggest that the signaling mechanism of LOV1 is noticeably different from that of LOV2. Compared with At phot, the photoreaction of Cr phot, excluding the photochemistry of the chromophore which has been studied, has not been explored fully. An FTIR study reported that light illumination of the LOV2 domain of Cr phot caused unfolding of the Jα helix 37, which is in accordance with the reaction of the LOV2 domain of At phot. A SAXS study showed that the molecular configuration of full-length Cr phot is changed slightly in the light state, which was explained by the separation of LOV2 from the kinase domain 38. This movement is tentatively attributed to the unfolding of the Jα helix. An MD simulation also reported that partial unfolding of the Jα helix of LOV2 is linked to kinase activation39. On the basis of these findings, the signaling mechanism of LOV2 appears to be rather well conserved between Cr and At phot. Indeed, the LOV2 domain of Cr phot predominantly regulates kinase activity and its flanking regions are also reported to be vital for signal transduction as observed for At phot 38. Moreover, Cr phot has been shown to rescue light responses in Arabidopsis when it is overexpressed, suggesting that the signaling mechanisms are conserved between higher plants and the green algae 40. On the other hand, however, there are some reports describing differences. The SAXS study reported that full-length Cr phot exists as a monomer in aqueous solution, indicating that LOV1 does not function to cause dimer formation

38

. Additionally, the MD simulation reported that the hinge region

predominantly forms a helical structure in the dark and the C-terminally flanking helix (hinge region) of LOV1 is unfolded in the light state 39. It was proposed that unfolding leads to changes of the molecular shape and the electrostatic character of the protein surface to bind its target molecules. These findings suggest that the photochemical properties of LOV1 of Cr phot differ from that of At phot; i.e., it is not a dimerization site but a light-sensing module. The recovery of the adduct form of LOV2 has been shown to be reduced ~2.5 times in the presence of LOV1, which might be responsible for enhancement of the light sensitivity of Cr phot because the photocycle speed in LOV2 is a key factor determining light sensitivity

38,41

. In the case of At phot, on the

other hand, LOV1 attenuates light sensitivity, which seems to be the opposite behavior to that of Cr phot 21. Surprisingly, the deceleration effect is retained even when the reaction of LOV1 is blocked, suggesting that alteration to light sensitivity is possibly achieved by static structural interactions between the LOV1 domain and other domains regardless of the photochemical state of LOV1 38. What is the biological relevance of the photoreaction of LOV1 in Cr phot? A yeast growth assay showed that inhibition of the photoreaction of LOV2 still shows weak light-dependent kinase activation 42, which indicates that the photoreaction of LOV1 is partially involved in signaling. In another study, overexpression of the N-terminal LOV domains (LOV1 + LOV2) was reported to affect eyespot size and 3



Photoreaction of Cr-Phototropin constructs phototaxis, suggesting that aside from activation of the kinase domain, these domains fulfill an independent signaling function in the cell

10

. Regulation was shown to be partially lost when the photoreaction of the

10

LOV1 domain is blocked . These findings suggest that the molecular basis of Cr-phot functioning would be different from that of At-phot, especially the photochemistry of the LOV1 domain. To provide deeper insight into the molecular mechanism of phot and into their similarity and/or diversity among different organisms, direct observation of photoreactions of LOV1 and LOV2 domains of Cr phot is required. In this study, we have investigated the reaction dynamics of LOV1, LOV1-hinge, LOV2, LOV2-linker and hinge-LOV2 (a construct containing LOV2 and its N-terminal extension) of Cr phot (Fig. 1(a)) by the TG method. We found that the reactions of these domains are very different from those of the corresponding domains of At phot. The differences and the diversity of phots are discussed. The variety of reactions found in this study should be important for understanding the reactions of the other LOV domains with the N- and C-terminal components. Furthermore, for understanding the reaction of full length Cr-Phot in future, the reactions of the constituted domains in this research will be useful and essential. EXPERIMENTAL SECTION Sample Preparation—Cr LOV1 (residues M1–S125), Cr LOV1-hinge (residues M1–I211), Cr LOV2 (residues T207–T321), Cr LOV2-linker (residues T207–K400) and Cr hinge-LOV2 (residues K126– T321) from Chlamydomonas reinhardtii phot cDNA were prepared using the polymerase chain reaction (PCR) with specific primer sets. The positions of truncations were determined by reference to a previous report.42 The recombinant proteins were overexpressed using Escherichia coli (E. coli) strain Rosetta 2 (Novagen) and the appropriate expression vector (pET-28a) with a His-tag at the N-terminus. The E. coli strains were grown at 37 °C in LB medium supplemented with kanamycin at 20 µg/mL until the culture reached an OD600 of 0.4–0.5 and protein overexpression was induced by the addition of isopropyl β-D-1-thiogalactopyranoside to a final concentration of 40 µM. The cells were incubated for a further 16 h at 18 °C. The harvested cells were suspended in a 50 mM Tris, 500 mM NaCl, pH 8.0 buffer containing a protease inhibitor cocktail (Nacalai tesque) and disrupted by sonication. The proteins were purified from cell free extracts with a HisTrap HP column (GE Healthcare), using a linear imidazole gradient (20 to 500 mM) in 500 mM NaCl plus 20 mM Tris, pH 8.0, to elute the protein from the HisTrap column. The eluted samples were further purified by size exclusion chromatography (Superdex 200 10/300 GL, GE Healthcare) and anion exchange chromatography (Mono Q, GE Healthcare) without cleavage of His-tag. After buffer exchange (10 mM NaH2PO4, 140 mM NaCl, 2.7 mM KCl, pH 7.4) using the Superdex 200 10/300 GL column, the purity of the proteins and chromophore occupancy were examined by SDS-PAGE and UV/Vis absorption spectra (Fig. 1(c) and (d)). The purity and the occupancy were > 95 % for all constructs. Blue light illumination induces the blue shift of absorption spectra for all samples (Fig. 1(c) and (d)), indicating that the adduct formation between the chromophore and the nearby cysteine residue is well conserved for all constructs examined in this study. Transient Grating Method—The experimental setup for the TG measurements was similar to that reported previously

26,32

. A laser pulse from a XeCl excimer laser (Lambda Physik, Compex102) -pumped 4


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Photoreaction of Cr-Phototropin constructs dye laser (Lumomics, HyperDye 300; 465 nm) was used as an excitation light source and a CW diode laser (Crysta Laser, 835 nm) as a probe light source. Sixteen signals were averaged by a digital oscilloscope (Tektronix, TDS-7104) to improve the signal-to-noise ratio. The repetition rate of excitation was usually 0.005 Hz. The q2 values at each experimental setup were determined from the decay rate of the thermal grating signal of the calorimetric reference (aqueous solution of bromocresol purple). All measurements were carried out at room temperature. Size-Exclusion Chromatography (SEC)—SEC measurements were performed using an ÄKTA purifier system with a Superdex 200 10/300 GL column (GE healthcare). The following size markers were used: 670 kDa, thyroglobulin (bovine); 158 kDa, γ-globulin (bovine); 44 kDa, ovalbumin (chicken); 17 kDa, myoglobin (horse); and 1.35 kDa, vitamin B12. The apparent molecular mass of the sample was determined from the calibration curve. The detection wavelength was 280 nm for all measurements. For the light-state experiments, continuous white light from a Xe lamp (MAX-300: Asahi Spectra) through a heat ray absorbing glass was illuminated on the column during the chromatographic experiments. The light intensity at the column position was about 60 mW/cm2and the temperature of the measurement was set to be 4 °C. Circular Dichroism Spectroscopy—Circular dichroism (CD) spectra were recorded with a spectropolarimeter (J720W1, JASCO, Osaka, Japan) with flowing N2 gas. The measurements were carried out in buffer solution (10 mM NaH2PO4, 14 mM NaCl, 0.27 mM KCl, pH 7.4). The background signal from the spectrum of the buffer solution was subtracted from all other measurements. The optical path length was 1.0 cm and the protein concentration was 0.5 µM for all samples. The measurements were performed with scan speed of 200 nm/min and 80 scans were averaged. To obtain the light-state CD spectra, the protein solution was illuminated with a blue LED (480 nm) for 1 min to accumulate the light state, and the illumination was stopped just before starting a scan. The temperature was set to be 4 °C to slow down the dark recovery rate. We repeated this scan 80 times and obtained spectra were averaged later. One scan took about 20 seconds, which was faster than the dark recoveries at the low temperature for all constructs. Since the light state recovered back to the ground state gradually during the recording, the obtained spectra might be slightly distorted. To measure the dark recovery process from the light adapted state more accurately at 20 °C, the samples were illuminated with the blue LED (480 nm) for 1 min and time traces of the changes in CD intensities at 220 nm were recorded. RESULTS Cr LOV1 Domain The TG signal of Cr LOV1 is shown in Fig. 2(a). The signal rose quickly within the response time of our system and showed a decay-rise-decay signal in the time range of 100 ns to 1 ms. Subsequently, the signal reached the baseline once and showed a rise-decay signal. The initial decay-rise and the subsequent decay component (< 1 ms of Fig. 2(a)) are expressed by a bi-exponential function (Equation 1) with rate constants of kf and Dthq2, ITG(t) = α{δnfexp(−kft) + δnthexp(−Dthq2t) + δnspe(t)}2

5


(Eq. 1)


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Photoreaction of Cr-Phototropin constructs where α is a constant representing the system sensitivity, and δnf, and δnth are the pre-exponential factors. The third term of Eq. (1) represents the time development in the slower time range, within 1 ms to 10 s. Because the qualitative profile in the faster time region (100 ns to 1 ms) is very similar to that observed for the photo-reactions of the LOV1 and LOV2 domains of At-phot 26,27,29,35, these phases were attributed to the adduct formation of the LOV1 domain and the thermal diffusion signal. The time constant of the adduct formation (kf−1) was determined to be 1.1 ± 0.1 µs, which is similar to a previously reported value for Cr LOV1 (800 ns (80%) and 4 µs (20%)) and that of other LOV domains (e.g. At-phot1LOV2 (1.9 µs) and At-phot2LOV2 (0.9 µs)) 26,27,43,44. The time range of the slower rise-decay signal was dependent on q2 (Fig. 2(b)). Therefore, we attributed the rise-decay component to the molecular diffusion process of the protein. The rise-decay profile (diffusion peak) indicates that the diffusion coefficient (D) of the product is different from that of the reactant. Because the refractive index change of the thermal grating signal (δnth) is negative under the present experimental conditions, we found that the signs of the rise and decay components were negative and positive, respectively. Hence, the rise and decay signals are assigned to the diffusion of the reactant and product, respectively, indicating that the photo-product diffuses slower than the reactant. The diffusion signal intensity was significantly dependent on q2 (Fig. 2(b)); i.e., the peak intensity was weak (or almost a single exponential decay) in a fast time range (large q2) and became stronger as the time range increased. This q2 dependence indicates that D of the photoexcited molecule is gradually changing over the observed time range. The observed TG signal during a wide observation time range (i.e., at various q2) was analyzed based on the following reaction scheme: hν k R → I → P

where R, I, P and k represent the reactant, an initial product (intermediate), the final product and the rate constant of the change, respectively. The time dependence of the refractive index changes due to the reactant (δnR(t)) and the product (δnP(t)) are given by

δnR (t) = δnR exp(−DRq2t)

δn P (t ) = δn I exp{ − ( D I q 2 + k )t} + δnP

k [exp{ − ( D I q 2 + k )t} − exp( − D P q 2 t )] 2 ( DP − DI ) q − k

(Eq. 2)

where DR, DI and DP are the diffusion coefficients of the reactant, intermediate and product, respectively. By global analysis, DR, DI and DP were determined to be (10.1 ± 0.2) × 10−11 m2 s−1, (10.1 ± 0.3) × 10−11 m2 s−1 and (7.6 ± 0.2) × 10−11 m2 s−1, respectively. The time constant of the D-change was determined to be 50 ± 10 ms at a concentration of 80 µM. The DR-value is similar to that of the monomeric form of At phot2LOV1 (9.8 × 10−11 m2 s−1, molecular mass: 17 kDa) and At phot2LOV2 (10.3 × 10−11 m2 s−1, molecular mass: 17 kDa)

26,45

. This

strongly indicates that the reactant is the monomer of LOV1 (molecular mass of the monomer: 16 kDa). According to the Stokes-Einstein relationship, under a given environment (T and η), D reflects the molecular size 46,47. If the difference in D between the reactant and the product (DR/DP = 1.33) is interpreted in terms of the difference in the molecular radius, the molecular volume of the product should be (1.33)3 = 2.34 times larger than that of the reactant. One of the possible explanations for the reduction of D is a dimerization 6


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Photoreaction of Cr-Phototropin constructs reaction of the monomeric LOV1 upon the photoreaction and this possibility is examined by measuring the concentration dependence on the diffusion signal and SEC profile of LOV1 in the dark and light states (Supporting Information: SI-1). The dimerization reaction of Cr LOV1 was indeed observed as a light induced shift of elution peak to higher molecular mass (Fig. S2(a) and (c)). Furthermore, we found that the diffusion signal intensity decreased with increasing concentration, and when the concentration was increased to 400 µM, a new rise-decay signal representing the dissociation of dimer into monomer appeared (Fig. S1(a)). This represents that Cr LOV1 exists as a dimeric form dominantly at high concentration (>400 µM) and it dissociates into monomer upon photoexcitation. Analyzing the temporal profiles obtained at high concentration (400 µM: Fig. S1(a) and (b)) by Eq. (2), the rate of dissociation was determined to be 60 s−1. Since the physiological concentration could be much lower than these concentrations, we do not discuss the dissociation reaction further in this main manuscript. The detailed analyses and discussions on the reactions of Cr LOV1 in a wide concentration range including the reaction scheme are described in SI-1.

Cr LOV1-Hinge The TG signal of Cr LOV1-hinge is depicted in Fig. 3(a). The profile in the short time scale region is very similar to that of Cr LOV1 described above. Thus, the assignment should be the same. The rate of the adduct formation was determined to be 0.8 µs, which is slightly faster than that of Cr LOV1 (1.1 µs), indicating that the hinge domain weakly affects the reaction rate of the chromophore. This change suggests that the hinge domain is located close to the chromophore. Based on the q2 dependence measurement, the next rise-decay signal was assigned to the diffusion signal. Interestingly, the intensity of the diffusion peak was much stronger than that of Cr LOV1. This stronger intensity indicates that the difference in D between the reactant and the product is larger for the Cr LOV1-hinge compared with that of Cr LOV1, as shown below. The diffusion signal intensity was strongly dependent on q2 (Fig. 3(b)); the signal intensity increased as the observation time range increased. We analyzed the time development of the diffusion signal (q2 dependence) by a similar method described in the above section and determined the following parameters: DR = (8.8 ± 0.3) × 10−11 m2 s−1, DI = (8.2 ± 0.4) × 10−11 m2 s−1, DP = (4.5 ± 0.2) × 10−11 m2 s−1, and the time constant of the D-change = 13 ms at 80 µM. Since DR is similar to that of the monomeric form of At phot2LOV1hinge (9.3 × 10−11 m2 s−1, molecular mass of the monomer: 26 kDa) and At phot1LOV2linker (9.2 × 10−11 m2 s−1, molecular mass of the monomer: 25 kDa) 26,27, the reactant should be a monomer of Cr LOV1-hinge (molecular mass of the monomer: 26 kDa). DR of Cr LOV1-hinge is smaller than that of Cr LOV1, which is expected because the molecular size of Cr LOV1-hinge is larger due to the presence of the hinge region. From the ratio of D between the reactant and product (DR/DP = 1.63), the volume change based on the Stokes-Einstein relationship was calculated to be (1.63)3 = 4.33, which cannot be explained solely by the dimerization reaction. This may indicate the formation of a higher oligomer and/or a change in the structure of the hinge region upon photoexcitation. To investigate the origin of the D-change, we measured the diffusion signal at various concentrations in different time ranges. Surprisingly, the concentration dependence on the diffusion signal in the slow time range was noticeably different from that of Cr LOV1. When the concentration was increased, 7



Photoreaction of Cr-Phototropin constructs the signal intensity was observed to increase slightly and it became almost identical above 160 µM (Fig. 4(a), q2 = 1.7 × 1010 m−2). The increase of the diffusion peak intensity contrasts the behavior observed for Cr LOV1 (Fig. S1(a)). This result can be explained by assuming that there is no equilibrium between the monomer and dimer in the dark, and only the reaction rate is dependent on the concentration. If the reaction is a multi-molecular process, the rate of the D change increases with increasing concentration. The diffusion peak intensity should also increase, because the number of D change events increases. At the higher concentrations (> 160 µM), the rate of the D-change became fast enough to complete the change before the diffusion signal, and consequently, the diffusion signal was not dependent on the concentration. Indeed, when we examined the concentration dependence at a larger q2, the concentration dependence became more apparent (Fig. 4(b)). Since the time scale of the diffusion signal overlapped with that of the reaction rate at this q2, the signal intensity was strongly affected by a change in the reaction rate. These results suggest that, in the case of Cr LOV1-hinge, there is no equilibrium between the dimer and monomer in the dark, and only the rate of the D-change is dependent on the concentration. Analyzing the signals at large q2 with Eq. (2) enabled determination of the rate constants at each concentration. Fig. 4(c) shows the obtained rate constants plotted against the concentration of Cr LOV1-hinge. The linear correlation shows that the reaction is a bimolecular reaction; i.e., dimerization. From the slope and intercept of the plot, the second order rate constant (kd) and the rate constant of the reverse reaction (krev) were determined to be (8.1 ± 1.2) × 105 M−1 s−1 and (41 ± 10) s−1. The oligomeric state of Cr LOV1-hinge was examined by SEC. We found that the elution peak position was not dependent on the concentration, which contrasts that observed for Cr LOV1 (Fig. S2(c) and (d)). The apparent molecular masses calculated from the peak elution times are 30 and 31 kDa for 40 and 400

µM, respectively, under dark conditions. Because the calculated molecular mass from the amino acid sequence of Cr LOV1-hinge is 26 kDa as the monomer, the SEC result implies that Cr LOV1-hinge exists as the monomer in the dark, which is consistent with the DR-value determined. Light illumination affected the absorption at 450 nm and the elution time significantly. The decrease of absorption at 450 nm indicates that enough amount of light state is accumulated under our experimental conditions (Fig. S2(b)). The apparent molecular mass of the light state determined by SEC (82 kDa) is much larger than the theoretical value of the dimer. This result indicates that Cr LOV1-hinge forms higher oligomeric species (e.g., trimers) due to additional interactions between residues in the hinge region and/or that the non-spherical shape of the dimer results in an overestimate of the molecular mass. Although we cannot exclude the first possibility, we presume that dimerization of the monomer is the dominant reaction of Cr LOV1-hinge, because the rate constant of the D-change was proportional to the concentration, indicating the bimolecular reaction (dimerization). Since DR/DP is very large, the conformation of the protein part is significantly changed upon dimerization. To investigate this postulate, we performed CD measurements to probe the secondary structural changes. The CD spectra of Cr LOV1 and Cr LOV1-hinge in the dark and light states are shown in Fig. 5(a). In the dark, the negative intensity at 208 and 222 nm, which predominantly represents the amount of the helical structure, was much stronger for Cr LOV1-hinge, indicating that the hinge region forms helices under dark conditions. Upon light illumination, shapes and intensities of CD spectra were changed significantly for 8


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Photoreaction of Cr-Phototropin constructs Cr LOV1-hinge, while Cr LOV1 showed minor changes. The dark recovery processes of the CD intensities at 220 nm of Cr LOV1 and Cr LOV1-hinge after blue light illumination are shown in Fig. 5(b). Importantly, a significant difference in the amplitude of the dark recovery between the two constructs was observed (190 ± 20 deg cm2 dmol−1 for Cr LOV1 and 390 ± 30 deg cm2 dmol−1 for Cr LOV1-hinge). The relatively small amplitude observed for Cr LOV1 indicates that the secondary structural changes are minor. Cr LOV1-hinge, however, showed a drastic change in the CD intensity when compared with that of Cr LOV1. We attribute this change to unfolding of the helical structure in the hinge domain in addition to movement of the LOV1 core. Hence, we conclude that Cr LOV1-hinge undergoes dimerization and unfolding of the helical structure upon light illumination, which leads to the observed larger decrease of D. The unfolding of the hinge helix may contribute to the overestimation of molecular mass (82 kDa) because of its non-spherical shape. If the dimerization and unfolding of the hinge helix occurs independently with different rates, the TG signals cannot be analyzed by Eq. (2), but by an equation containing two rate constants. The fact that the signals were consistently reproduced well by one rate constant indicates that dimerization and unfolding occur simultaneously. Additionally, if the unfolding of the hinge helix is a rate-determining step of dimerization, the rate of the D-change should not depend on the concentration. The clear concentration dependence on the rate indicates that dimerization is the rate-determining step, that is, the hinge helix is destabilized upon dimerization and unfolds immediately. There are two possible reaction schemes for light-induced dimer formation: the ground state monomer and excited monomer form the dimer, or two excited monomers form the dimer. To distinguish these, we measured the diffusion signal at various excitation light intensities over a range of 3.3–260

µJ/pulse (Fig. S5(a)) and found that the square root of the TG signal intensity, which is proportional to the number of dimerization events, linearly increased with the laser power intensity (Fig. S5(b)). This observation shows that dimerization occurs between the Cr LOV1-hinge in the light state and in the dark state. When the power increased above 30 µJ/pulse, however, the laser power dependence is saturated. At the same time, when we plotted the square root of the TG signal intensity before the diffusion signal, representing the number of adduct species, it shows similar behavior (Fig. S5(c)). The laser power dependence was fitted by the following equation representing the saturation effect of light absorption,

αn = α /(1+ I / IS)

(Eq. 3)

0

where αn is the measured non-linear absorption coefficient, α0 is the absorption coefficient at a low light intensity, I is the laser intensity, and IS is the saturation intensity. The numbers involved in the adduct formation and that in the subsequent dimerization were well reproduced by Eq. 3 with the same IS (33

µJ/pulse). This result indicates that the number of the dimerization events is proportional to that of the adduct formation. Since the dimeric form of Cr LOV1-hinge was dominant under a continuous blue-light illumination condition under which most of Cr LOV1-hinge was converted to the light state (Fig. S2(b) and (d)), it might be reasonable to consider that two excited monomers also formed the dimer. However, the present measurement indicates that one photoexcited monomer is sufficient for dimer formation. The question that arises at this point is whether both monomer units in the dimer undergo unfolding of hinge helix or does only one monomer unit in the dimer show unfolding? To answer this question, we 9



Page 10 of 37

Photoreaction of Cr-Phototropin constructs compared the temporal profiles of the diffusion signals obtained under various laser power intensities. Fig. S5(d) shows the signals normalized with the peak intensity. Interestingly, they superimpose well even though the laser intensity was high enough to saturate the photoexcitation process. This result indicates that the D value of the dimer containing one excited monomer and one ground state monomer (LOV1*-LOV1) is the same as that of a dimer containing two excited monomers (LOV1*-LOV1*). If the unfolding of the hinge helix occurs only for the excited molecule, the dimer of LOV1*-LOV1 should possess a different D value from that of LOV1*-LOV1*. The superposition of the diffusion signals indicates that the D-change does not depend on the extent of excitation, and hence we conclude that the secondary structural changes observed by the CD measurements should occur in the dimer of LOV1*-LOV1 and LOV1*-LOV1*. Therefore, this conformational change is induced by the dimerization reaction and not by formation of the adduct.

Cr LOV2 and Cr LOV2-Linker The TG signals of Cr LOV2 and Cr LOV2-linker are depicted in Fig. 6(a). In the fast time scale, the figure shows that adduct formation and the thermal diffusion signals are similar to other LOV domains. The rate of adduct formation was determined to be 0.2 µs for all samples. The same rate constants indicate that the linker regions do not affect adduct formation. Signals appearing in the slow time scale (~ 1 s) were assigned to the molecular diffusion signals for all constructs. Interestingly, Cr LOV2 showed almost monotonous decay signals, whereas Cr LOV2-linker showed the rise-decay profile. The monotonous decay indicates that the photoexcitation of Cr LOV2 does not lead to significant D-changes. When the full-linker was included, however, the D-change was observed, indicating that the linker domain is relevant for the D-change. The decay signal of Cr LOV2 can be reproduced well by

ITG(t) = α{(δnP − δnR)exp(−DRq2t)}2

(Eq. 4).

The D value (DR = DP) was determined to be (9.7 ± 0.1) × 10−11 m2 s−1. This value is similar to those of the monomer of the LOV domains, indicating that Cr LOV2 exists as a monomer in solution both in the dark and light states, and it does not show any diffusion sensitive conformational change. The rise and decay components of the diffusion signal of Cr LOV2-linker were assigned to the diffusion of the reactant and the product, respectively (DP < DR). Since only Cr LOV2-linker showed the rise-decay signal representing the significant D change upon photoexcitation, we analyzed the time development of the diffusion signal for this construct. The diffusion signal intensity was dependent on q2, as shown in Fig. 6(b); the signal decayed monotonously at high q2 and it showed a diffusion peak at relatively low q2 conditions. The diffusion signal intensity increased as the observation time range increased. We analyzed the time development of the diffusion signal in a similar way, as described in the above sections (Eq. (2)), and determined the following parameters: DR = (8.3 ± 0.3) × 10−11 m2 s−1, DI = (8.3 ± 0.3) × 10−11 m2 s−1, DP = (6.5 ± 0.2) × 10−11 m2 s−1 and the time constant of D-change = 1.3 s at 200 µM. The DR value implicates that Cr LOV2-linker exists as a monomer in the dark state. From the ratio of the D values between the reactant and product (DR/DP = 1.28), the volume change was calculated to be (1.28)3 = 2.10 using the Stokes-Einstein relationship. This number may indicate that a possible origin of the D-change is the dimerization reaction. 10


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Photoreaction of Cr-Phototropin constructs To investigate the origin further, we measured the diffusion signal at various protein concentrations. Fig. S6(a) shows the concentration dependence on the diffusion signal at q2 = 1.3 × 1010 m−2. When the concentration was increased, the signal intensity increased, even though the signals were normalized by the number of photo-adduct species. This result indicates that the rate of D-change increases with increasing concentration. The rate constants were determined at each concentration by analyzing the signals using Eq. 2. We found that the rate linearly correlates to the concentration (Fig. S6(b)), which is consistent with the dimerization reaction. From the slope and intercept of the plot, the kd and krev were determined to be (2.9 ± 0.7) × 103 M−1 s−1 and (0.18 ± 0.08) s−1, respectively. The oligomeric state was examined by SEC measurements and the results are shown in Fig. S7. The peak positions of Cr LOV2 did not depend on the light condition and the molecular mass was determined to be 20 kDa for Cr LOV2 (Fig. S7(a) and (c)). This corresponds to the monomer of Cr LOV2 (15 kDa). In the case of Cr LOV2-linker, the peak shifted to a higher molecular mass upon light illumination and the molecular masses determined from the peak positions were 22 kDa for the dark state and 33 kDa for the light-adapted state (Fig. S7(b) and (d)). The molecular mass for the dark state is close to that of the monomer of Cr LOV2-linker (23 kDa). The increase in the molecular mass upon light illumination clearly indicates that Cr LOV2-linker associates to form the dimer in the light state. The smaller value compared to the calculated value for the dimer may be explained by an insufficient accumulation of the light state due to its relatively fast dark recovery rate, as shown later. The single peak of the elution profile indicates an equilibrium between dark and light state during the elution (exchange between monomer and dimer), which is fast enough compared with that of the elution time. Due to this fast equilibrium, the monomer and dimer peaks do not give rise to two separate peaks but one broadened elution peak as observed in Fig. S7(d). Regarding the concentration dependence on the elution profile, all samples did not show any shift of the peak position upon changing the initial concentration. Hence, we conclude that Cr LOV2 exists as a monomer, whereas light illumination induces dimerization of Cr LOV2-linker. Previous studies have shown that the C-terminal region of At LOV2 forms a helical structure in the dark and it fully or partially unfolds in the light state, and these reactions are detected as significant decreases of D by the TG method 26,27,29,32. However, our TG measurements found that Cr LOV2-linker dimerized, and its D-change is explained by only a volume change associated with dimerization. To examine the secondary structure change by light irradiation, CD spectra were measured (Fig. 7(a)). The negative intensity at 208 and 222 nm was increased as the linker domain is included, which clearly represented that linker region forms the helical structure in the dark. However, both samples showed very similar amplitude changes associated with the dark recovery (Fig. 7(b)) (220 ± 20 deg cm2 dmol−1 for LOV2 and 190 ± 10 deg cm2 dmol−1 for LOV2-linker). Hence, we consider that the secondary structure change occurs not in the linker helices but in the LOV2 core. Previous studies on the LOV domain of YtvA, which is a blue light sensor in Bacillus subtilis, have shown similar changes in the CD spectrum upon light illumination, which was attributed to a distortion of the central β-scaffold of the LOV domain 48,49. Therefore, the CD intensity change observed for Cr LOV1 is possibly attributed to movement of the β-strands. These results are different from what we observed for LOV2-linker of At phot, in which the linker helix unfolds in the light state 32.

11




Hinge-LOV2 In the above section, we showed that the hinge domain changed drastically in secondary structure upon photoexcitation of the LOV1 domain. This result was not expected based on the previous studies of At LOV1-hinge. Consequently, is unfolding of the hinge helix triggered only by the LOV1 domain? In other words, can LOV2 also regulate the conformation of the hinge domain that locates to the N-terminal side? To investigate this question, we measured the TG signals of Cr hinge-LOV2. Interestingly, the molecular diffusion signal showed a rise-decay profile (Fig. S8(a)), which is different from that of Cr LOV2, which shows almost monotonous decay signal. This shows that the hinge domain is involved in the D-change upon photoexcitation of LOV2, as observed for LOV1. From the signs of refractive index changes, the rise and decay components are assigned to the diffusion of the reactant and product, respectively, i.e., DR > DP. We observed the time development of the diffusion signal as shown in Fig. S8(b). Analyzing the data by Eq. 2 yielded DR, DI, DP and k−1 values of (7.2 ± 0.3) × 10−11 m2 s−1, (7.0 ± 0.4) × 10−11 m2 s−1, (5.4 ± 0.3) × 10−11 m2 s−1 and 2.9 ± 1.5 s, respectively. The DR value suggests that hinge-LOV2 exists as a dimer in the dark. The magnitude of the D change (DR/DP = 1.33) is relatively minor compared with that of LOV1-hinge (DR/DP = 1.63), which undergoes dimerization and unfolding of the hinge helix. To reveal the origin of the D-change, we measured the concentration dependence of the diffusion signal (Fig. S9). The detailed analyses are described in Section SI-5 (Supplemental Data). On the basis of these analyses, we concluded that the dimerization reaction is the main reaction upon photoexcitation of this sample. The CD spectrum of Cr hinge-LOV2 in the dark state is shown in Fig. 7(a). The negative amplitude at 208 nm and 222 nm increased when the hinge domain was present, indicating that the hinge region forms a helix. Comparison of the difference in the CD spectra due to the presence of the hinge domain in LOV1 and LOV2 showed that the hinge domain present in LOV1 has a larger impact on the ellipticity values (Figs. 5(a) and 7(a)). Here, increases in the CD intensities at 222 nm by the presence of the hinge domain were 810 deg cm2 dmol−1 for LOV1 and 420 deg cm2 dmol−1 for LOV2. This shows that the amount of helical structure in the hinge domain is smaller for the hinge-LOV2 than that of LOV1-hinge, indicating that a part of the hinge helix is unfolded in the hinge-LOV2 even in the dark state. The dark recovery of the CD intensity of Cr hinge-LOV2 is shown in Fig. 7(b). The amplitude of the dark recovery of the CD signal (210 ±30 deg cm2 dmol−1 for Cr hinge-LOV2) was similar to those of other LOV2 constructs. This indicates the unfolding of the hinge helix is not induced by excitation of LOV2. From these observations, we conclude that the D-change is attributed not to the unfolding of the hinge helix but to dimerization. A part of the hinge helix is unfolded in the hinge-LOV2 (i.e., without the LOV1 domain) even in the dark state.

Dark Recovery of LOV1 and LOV2 Previous studies suggested that the rates of dark recovery of the LOV domains are related to their light sensitivity; i.e., the slower the recovery is, the higher the sensitivity is. LOV1 is hypothesized to regulate the light sensitivity of phot, which is supposed to be achieved by controlling the dark recovery rate of LOV2 38,41. Indeed, it has been reported that the dark recovery rate of Cr LOV2 is reduced in the presence of LOV1 38. Here, we studied the effect of the flanking domains of LOV1 and LOV2 on the lifetime of the 12


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Photoreaction of Cr-Phototropin constructs light adapted form by monitoring the absorption change associated with the dark recovery process. Fig. 1(c) and (d) show the absorption spectra of constructs used in this study in both dark and light states. The spectra are not affected by the presence/absence of the flanking region of the LOV1 and LOV2 domains. Fig. S11(a) and (b) show the dark recoveries of absorption at 450 nm. In the case of LOV1, the signal was reproduced by a single exponential function and the dark recovery rate was hardly affected by the presence of the hinge domain (118 ± 3 s for Cr LOV1, 125 ± 8 s for Cr LOV1-hinge). This indicates that the hinge is not involved in the stabilization of the light state. Interestingly, however, the dark recovery of LOV2 was bi-exponential and the rates were significantly affected by the presence of flanking regions (23 ± 2 s (49%) and 96 ± 4 s (51%) for Cr LOV2, 10 ± 1s (67%) and 19 ± 1 s (33%) for Cr LOV2-linker, 35 s ± 10 (3%) and 290 ± 60 s (97%) for Cr hinge-LOV2). In particular, when the hinge domain was included, the recovery rate was reduced significantly. This indicates that the hinge domain is located close to the chromophore of LOV2 and stabilizes the adduct form to slow down the recovery process dramatically. On the basis of these results, we propose that the hinge domain, and not the LOV1 domain, is a dominant regulator of the recovery rate of Cr LOV2. In addition, the dark recoveries monitored by the CD intensities at 222 nm were also analyzed and the lifetimes of the light states are listed in Table S1. Mostly, adduct-state lifetimes from the CD-signals and from the absorptions agree well. Although we found that the recovery lifetimes from the absorption signals are slightly shorter than those from the CD measurements for LOV1, LOV1-hinge, LOV2, LOV2-linker, this difference may be explained by the effect of the probe light. For the absorption measurement, the sample solution should be exposed to the blue light, which converts the dark state to the light state. Though we used a neutral density filter in front of the sample solution to reduce the intensity and minimize the effect of the probe light, the probe light may still accelerate the apparent rate constant, which is a sum of the rate constants of the (weak) excitation and of the dark recovery. Hinge-LOV2 is exceptional. The lifetime from the absorption change signal is slower than that obtained from the CD measurement. The origin of this difference is not clear at present. The obtained parameters on the photoreactions of all constructs examined in this study are listed in Table. 1. The parameters of At phot2 obtained from previous studies are also listed for comparison 26,29.

DISCUSSION Light induced conformational changes of LOV1 constructs We found that Cr LOV1 is in dynamic equilibrium between the monomer and dimer states. In a relatively low concentration range, the monomer is dominant, and the light-induced dimerization is the main reaction (Fig. 8(a)). A previous study using a crosslinking technique on Cr LOV1 also showed two bands corresponding to the monomer and dimer, and the dimer population was found to increase significantly upon light illumination, and relax back to the initial ratio during incubation in the dark 50. The observation of the light-dependent association at low concentrations is consistent with our findings. Although it was found that Cr LOV1 is in equilibrium between the monomer and dimer, Cr LOV1-hinge exists as a monomer in the dark, indicating that the dimerization site of LOV1 in the dark is masked by the hinge domain. Since the β-strands of the LOV1 core are probably the dimerization site as 13



Photoreaction of Cr-Phototropin constructs suggested by the crystal structure, the hinge region may dock against the β-strands in the dark. Upon photoexcitation, Cr LOV1-hinge undergoes dimerization and the unfolding of the hinge helices (Fig. 8(b)). Since dimerization is the rate-determining step for the unfolding of the hinge domain, the helical structure of the hinge should be destabilized by the dimerization. This dimerization-then-unfolding scheme is consistent with the unfolding of the hinge helix of not only an excited monomer but also another (ground state) monomer unit in the dimer. These findings suggest that the dimerization site in the light state is different from that in the ground state. Using the crystal structure of Cr LOV1, we performed a docking simulation to predict the homodimer structures for (LOV1-LOV1) and (LOV1*-LOV1) and the highest ranked dimer structures are shown in Fig.S4. It is interesting to note that these structures are different. Although we cannot evaluate the adequacy of these simulated structures, we consider that the dimerization site is very sensitive to the minor structure change. The determined intrinsic dimerization rate constant (kd = 8.1 × 105 M−1 s−1) was much smaller than that of a diffusion-controlled reaction calculated by the Smoluchowski−Einstein equation for a bimolecular reaction in solution (~109 M−1s−1)

51

. This difference indicates that the collision between two protein

molecules is not sufficient for dimerization, but their relative orientations dictates additional constraints, which reduces the rate of the reaction by three orders of magnitude. Furthermore, this small kd suggests a very small steric factor; that is, the dimerization reaction occurs only at a specific location of the protein. The rate of dimerization of Cr LOV1 was determined only at the low concentration (80 µM) and the value was 50 ms. At the same concentration, the dimerization rate of Cr LOV1-hinge was determined to be 13 ms which is faster than that of Cr LOV1, indicating that the steric factor is smaller for Cr LOV1 than that of Cr LOV1-hinge. By considering that the hinge covers the dimerization site of LOV1 in the dark, the dimerization reaction between excited monomer and ground state monomer of LOV1-hinge should be reduced dramatically if light induced dimerization occurs at the same site. The relatively faster rate of the dimerization of LOV1-hinge supports the idea that dimerization occurs at a different site from that of the ground state dimer of Cr LOV1 (β-strands), as discussed above. What is the biological relevance of the light induced dimerization of LOV1? There are several reports implying that the LOV1 domain regulates the biological function in a light-dependent manner. Recently, it has been reported that a construct containing only the N-terminal region of Cr phot (LOV1+LOV2) regulates eyespot size and phototaxis of Chlamydomonas reinhardtii, suggesting that the light-sensing domains fulfill an independent signaling function in the cell aside from activation of the kinase domain

10

. This might be achieved by an interaction with other proteins and it has been shown that

inactivation of the LOV1 domain (C57S) partially loses regulation 10. If the signaling of the LOV domains have multiple pathways via interacting with other proteins in the cell, the light induced dimerization of the LOV1 domain and structural change to the hinge region observed in this study are possibly involved in these additional processes. A SAXS study on full-length Cr phot has shown that the LOV1, LOV2 and kinase domains are in a collinear arrangement

38

. The molecular configuration shows a slight bend in the middle upon light

illumination, which has been explained by the slight separation of the LOV2 domain from the kinase domain due to conformational changes in the flanking region of the LOV2 domain 38,41. On the basis of our findings, 14


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Photoreaction of Cr-Phototropin constructs we consider that the change in the overall shape should be at least partially attributed to the conformational change to the hinge domain and the reorientation between LOV1 and LOV2. Since the oligomeric structure of full-length Cr phot has been reported to be a monomer both in the dark and light states, there is a possibility that the homo-dimerization of the LOV1 domain observed here corresponds to the interaction with another domain in the intact protein.

Conformation changes of the linker We found that the oligomeric states of the LOV2 domains are strongly dependent on the length of the C-terminal extension (Fig. 9(a–c)). In the dark, LOV2 and LOV2-linker exist as a monomer. Upon light illumination, only LOV2-linker undergoes dimerization, which is presumably caused by the exposure of the dimerization site at the linker region due to a structural change to the linker domain. The rate constant (kd = 2.9 × 103 M−1 s−1) was much smaller than a calculated diffusion-controlled reaction rate and even smaller than that of Cr LOV1-hinge. This small kd suggests a very small steric factor for the dimerization of LOV2-linker. This result suggests that the dimerization site is different between LOV1 and LOV2. In contrast to the case of LOV1-hinge, the secondary structure of the C-terminal region of the LOV2 domain was silent to light illumination. Although the CD measurement detected that the linker domain forms helical structures predominantly in the dark, changes in CD intensity upon light illumination were well explained not by unfolding of the helices but by conformational changes of the LOV2 core (distortion of β-strands). The TG measurement also showed that the D-change is minor for LOV2-linker, and it is well explained by only the dimerization reaction, indicating that the diffusion sensitive conformational change is not induced in the linker domain. In a previous study, unfolding of the helical structure of Jα has been reported for Cr LOV2 by FTIR 37. However, we consider that these results are not inconsistent, because physical properties for these studies are different. The vibrational data is sensitive to rather local changes of the structure; e.g., even a small change can induce the vibrational frequency change. On the other hand, the diffusion change is not sensitive to a small local conformation change, but reflects a rather large change that affects the solvent-solute interaction. Hence, it is not surprising that a minor structural change was not apparent in the TG signal. Indeed, an MD simulation has shown that the unfolding of the Jα helix of Cr LOV2 is relatively minor compared with that of the C-terminal helix of the LOV1 domain

39

. The MD

simulation also suggested that the slight unfolding of Jα is sufficient to ensure entry of an ATP molecule into the catalytic site of the kinase domain. In the case of phot from higher plants, light induced unfolding of the Jα helix has been reported extensively 22–24,52. Our recent work on At phot2 has also clarified that the linker domain outside the Jα helix also undergoes unfolding, which seems to be related to kinase activation

29

. Utilizing these dynamic

structural changes, higher plants may achieve more robust signal transduction when compared with that of Cr phot. Cr phot has been reported to rescue phot function in At only when it is overexpressed

40

, which

indicates that the efficiency of kinase activation would be lower for Cr phot due to the minor conformation change of the linker domain. What creates diversity in the photoreaction among different organisms? The Jα helix of Avena sativa (As) phot1LOV2 has been reported to display an amphipathic character, showing characteristic 15



Page 16 of 37

Photoreaction of Cr-Phototropin constructs 53

hydrophobic and polar faces on the helix . Additionally, the interface between the LOV2 core and the Jα helix is composed of hydrophobic residues, and this interaction may stabilize the helical structure of Jα. When a charged residue is introduced into the hydrophobic side of the Jα helix, the interaction is disrupted and the helix is disordered

53

. The amphipathic character is well conserved in At phot1LOV2 and At

phot2LOV2, and they show unfolding of the Jα helix upon light illumination

26,27,29

. In case of Cr phot, on

the other hand, the Jα region does not possess amphipathic character (Fig. S12) and Cr LOV2-linker did not unfold upon photoexcitation. This indicates that the helical structure may be stabilized not by hydrophobic interactions with the LOV2 core but by another factor. In such a case, the dissociation of the Jα helix from the LOV2 core does not destabilize the helical structure, and subsequently, unfolding is not induced. We speculate that the amphipathic character of the Jα helix is a key feature for reactivity.

Function of hinge in Cr phot In phots, the output domain (kinase) is located on the C-terminal side of the photoreceptor LOV domain and the signal proceeds through conformational changes to the linker domain. However, in the case of Cr-phot, the conformational change of the linker is minor and an alternative signaling pathway may exist. Another LOV protein aureochrome is a unique LOV-photoreceptor, because the signal output domain exists on the N-terminal side of the LOV domain 54,55, which indicates that the LOV domain is able to transmit the light signal not only to the C-terminal side but also to the N-terminal side. Indeed, recent studies on phot have reported that the A'α-helix in the N-terminal upstream LOV2 plays an important role in conformational changes and signal transduction

28,42,56

. In the case of Cr-phot, the hinge domain shows significant

conformational change upon photoexcitation of the LOV1 domain, which indicates that the hinge domain shows high reactivity. Therefore, we investigated the photoreaction of hinge-LOV2 to study the signaling direction of the LOV2 domain. However, we found that the hinge domain was already unfolded partially in the dark state and we could not determine if the LOV2 domain induces structural changes to the hinge domain in a light-dependent manner. These results strongly suggest that the helical structure of the hinge domain is stabilized by interaction with the LOV1 domain. This indicates that the photoexcitation of LOV1 leads to dimerization and this may result in the dissociation of the hinge domain, and destabilization of the helical structure. Previous studies suggested that the lifetime of the light state of the LOV2 domain is related to the light sensitivity of phot

38,41,57–59

. The presence/absence of the LOV1 domain affects the recovery rate of

LOV2 and this is a function of the LOV1 domain, that is, controlling the light sensitivity of phot 38. However, it has been reported that the photochemical state of LOV1 is not important for the control of the lifetime. This casts a question about the role of LOV1, i.e., how does LOV1 control light sensitivity? Light sensitivity should remain constant, because LOV1 always exists in the intact protein. In this study, we found that the recovery rate of LOV2 is strongly affected by the presence of its N- and C-terminal extensions, while the recovery rate of LOV1 is not sensitive to the presence of its C-terminal region. In particular, the presence of the hinge domain strongly slows down the recovery rate of LOV2 indicating that not the LOV1 but the hinge is a main factor for controlling the dark recovery rate of LOV2. If this is the case, the light-independent

16


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Photoreaction of Cr-Phototropin constructs deceleration effect is explained reasonably, because the hinge domain always locates to the N-terminal side of the LOV2 domain. The biological role of the LOV1 domain requires further validation.

Comparison with other LOV domains So far, many proteins that contain the LOV domain have been found. Such LOV proteins exist as single- and multi-domain proteins, and they transmit the light-induced signal to downstream protein modules via intermolecular and/or intramolecular interactions. Current many researches indicate that, in nearly all cases, the LOV domain signals are transmitted to the effector elements through highly variable N-terminal or C-terminal extensions to the LOV core. For example, the unfolding of the C-terminal helices of the LOV2 domains are vital steps for signaling of Avena sativa (As) and At phots.20,22,53 YtvA, which consists of a LOV domain and a STAS domain regulating stress response in Bacillus subtilis, exists as a dimer in both dark and light states and the C-terminal Jα helix extends from the LOV core dimer in a coiled-coil arrangement.60,61 Upon light illumination, a scissor-like rotational motion of the C-terminal helices in the dimer has been proposed to be relevant for signal transduction. A similar rotational motion of the Jα helix was also reported for a bacterial short LOV protein PpSB1.62,63 Another LOV protein VVD, which is a short LOV protein from Neurospora crassa containing only a LOV domain and its N-terminal extension, exposes a hydrophobic surface, which supports homodimer formation through reorganization of the N-terminal elements upon photoexcitation.64,65 Aureochrome1, which is a blue-light regulated transcriptional factor from Vaucheria frigida, also forms the dimer in the light state.66 The photoinduced dimerization is mediated by the LOV domain dominantly, which is relevant for the following DNA binding. During the dimerization process, the flanking N- and C-terminal helices of the LOV domain are considered to undergo conformational changes.67 These established models for the molecular mechanisms of the LOV-mediated signal transduction exhibits the diversity among a variety of LOV proteins. In this study, we also found that the photochemistry of the LOV domains and its flanking regions are different between Cr phot and At phot as summarized in Table 1, which was not apparent before this study. Even though the initial adduct formation reaction and oligomeric structures of the LOV domains are well conserved between Cr phot and At phot, the later reactions of the constructs containing N- and C-terminal extensions exhibit different characters. These findings indicate the important role of the flanking regions to control the inter- and intra-molecular interactions of the LOV proteins. Dimeric structure is also an important factor for describing the LOV domain and also for understanding the reactions. The β-scaffold of the LOV core is employed for the dimerization in At phot1 LOV1 and At phot2 LOV1.34 However, in a case of At phot1 LOV2, the N-terminal helix largely contributes to the dimerization through an α-helical coiled coil.30 The N-cap mediated dimer interface is also reported for VVD and the LOV protein from the marine bacterium Dinoroseobacter shibae (DsLOV).61,65,68 The structures of YtvA and PpSB1 show that the dimer interfaces are mediated by both the N- and C-terminal auxiliary helices. A dimer structure of the LOV protein from Rhodobacter sphaeroides (RsLOV) exhibits the N- and C-terminal helical extensions that form a helical bundle at its dimer interface.69 On the basis of these findings, it is clear that the interface of the dimer structure shows diversity among the LOV proteins, and a slight difference in the structure of the LOV core and N- and C-terminal extensions easily affect the stability 17



Photoreaction of Cr-Phototropin constructs and structure of the dimeric form. The high sensitivity of the dimer structure to the LOV domain structure could be an important origin to the variety of reactions of the LOV proteins. The different dimer interface between the dark and light adapted dimers found in this research may reflect this high sensitivity. It may not be surprising to find that the dark recovery rate from the light state of the LOV domain depends on residues within the LOV domain. In this research, we found that the dark recovery rate is strongly dependent on the presence/absence of N- and C-terminal extensions for Cr LOV2. This sensitivity to the dark recovery rate was not reported for the LOV domain reactions of At phot. However, bacterial short LOV proteins PpSB1 and PpSB2 display drastically different dark recovery rates (~ 40 h for PpSB1 and ~140 s for PpSB2), though both proteins are highly similar in sequence (66% identity).70 When their C-terminal extensions (Jα helices) are swapped with each other, the recovery rates are accelerated and decelerated significantly for PpSB1 and PpSB2, respectively. Hence, this is another diversity showing that the recovery rates of the LOV domains are sensitive to not only the residues within the LOV domain, but also the outer domains.

CONCLUSIONS In this study, we investigated the reaction dynamics of the LOV domains and their flanking regions of Cr phot using the TG method. We clarified that the Cr LOV1-hinge shows drastic structural changes in the hinge domain in addition to the dimerization upon the photoexcitation. These reactions should be related to its function. Although it has been proposed that the LOV1 domain controls the light sensitivity of phot through changing the dark recovery rate of the LOV2 domain, we found that the presence of the hinge domain is sufficient to slow down the dark recovery of the LOV2 domain, indicating that LOV1 might not be involved in controlling light sensitivity. In the case of Cr LOV2-linker, we found that dimerization is the main reaction and structural changes of the linker domain are minor. Since previous biochemical studies reported that the LOV2 of Cr phot is a main regulator of the kinase activity in a light-dependent manner, we suggest that the minor structural change is sufficient to activate the kinase domain as suggested by an MD simulation study 39. Interestingly, we found that the photochemical properties of Cr phot are considerably different from those of At phot regarding conformational changes (D-changes) and their kinetics (Table 1), although Cr phot has been reported to rescue the phot function in At. This indicates that the molecular mechanisms of signal transduction are not necessarily the same even though the biological response is similar. It suggests that the molecular mechanism of the function have changed during the course of the evolution. The conformational change induced by LOV1 might be unnecessary and it has been reduced in higher plants, while the drastic conformational change of the linker domain is required for robust signal transduction and higher plants have acquired the property. Understanding what gives the diversity to the photoreaction at the molecular level is important. Since we now hypothesize that the amphipathic character of the linker domain is a key factor determining the reactivity, it will be interesting to swap the linker domains between At phot and Cr phot and detect their photoreactions.

SUPPORTING INFORMATION 18


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Photoreaction of Cr-Phototropin constructs The Supporting Information is available free of charge on the ACS Publications website. SI-1; Reaction dynamics of Cr LOV1, SI-2; Laser power dependence on TG signal of LOV1-hinge, SI-3; Concentration dependence of diffusion signal of LOV2-linker, SI-4; SEC profiles of Cr LOV2 and Cr LOV2-linker, SI-5; Reaction dynamics of Cr hinge-LOV2, SI-6; The dark recovery of adduct form, SI-7; Helical wheel analyses on linker region of Cr phot and At phot1.

ACKNOWLEDGEMENTS: This work was supported by a Grant-in-aid for Scientific Research on Innovative Areas (research in a proposed research area) (Nos. JP20107003, and JP25102004) and a Grant-in-aid for Scientific Research (25288005, 17H03008) from MEXT/JSPS (to M.T.).

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REFERENCES (1)

Christie, J. M. Phototropin Blue-Light Receptors. Annu. Rev. Plant Biol. 2007, 58 (1), 21–45.

(2)

Christie, J. M.; Reymond, P.; Powell, G. K.; Bernasconi, P.; Raibekas, A. A.; Liscum, E.; Briggs, W. R. Arabidopsis NPH1: A Flavoprotein with the Properties of a Photoreceptor for Phototropism. Science 1998, 282 (5394), 1698–1701.

(3)

Kagawa, T.; Sakai, T.; Suetsugu, N.; Oikawa, K.; Ishiguro, S.; Kato, T.; Tabata, S.; Okada, K.; Wada, M. Arabidopsis NPL1: A Phototropin Homolog Controlling the Chloroplast High-Light Avoidance Response. Science. 2001, 291 (5511), 2138–2141.

(4)

Jarillo, J. A.; Gabrys, H.; Capel, J.; Alonso, J. M.; Ecker, J. R.; Cashmore, A. R. Phototropin-Related NPL1 Controls Chloroplast Relocation Induced by Blue Light. Nature 2001, 410 (6831), 952–954.

(5)

Kinoshita, T.; Doi, M.; Suetsugu, N.; Kagawa, T.; Wada, M.; Shimazaki, K. phot1 and phot2 Mediate Blue Light Regulation of Stomatal Opening. Nature 2001, 414 (6864), 656–660.

(6)

de Carbonnel, M.; Davis, P.; Roelfsema, M. R. G.; Inoue, S.-I.; Schepens, I.; Lariguet, P.; Geisler, M.; Shimazaki, K. I.; Hangarter, R.; Fankhauser, C. The Arabidopsis PHYTOCHROME KINASE SUBSTRATE2 Protein Is a Phototropin Signaling Element That Regulates Leaf Flattening and Leaf Positioning. Plant Physiol. 2010, 152 (3), 1391–1405.

(7)

Huang, K.; Merkle, T.; Beck, C. F. Isolation and Characterization of a Chlamydomonas Gene That Encodes a Putative Blue-Light Photoreceptor of the Phototropin Family. Physiol. Plant. 2002, 115 (4), 613–622.

(8)

Huang, K.; Beck, C. F. Phototropin Is the Blue-Light Receptor That Controls Multiple Steps in the Sexual Life Cycle of the Green Alga Chlamydomonas Reinhardtii. Proc. Natl. Acad. Sci. U. S. A.

2003, 100 (10), 6269–6274. (9)

Im, C. S.; Eberhard, S.; Huang, K.; Beck, C. F.; Grossman, A. R. Phototropin Involvement in the Expression of Genes Encoding Chlorophyll and Carotenoid Biosynthesis Enzymes and LHC Apoproteins in Chlamydomonas Reinhardtii. Plant J. 2006, 48 (1), 1–16.

(10)

Trippens, J.; Greiner, A.; Schellwat, J.; Neukam, M.; Rottmann, T.; Lu, Y.; Kateriya, S.; Hegemann, P.; Kreimer, G. Phototropin Influence on Eyespot Development and Regulation of Phototactic Behavior in Chlamydomonas Reinhardtii. Plant Cell 2012, 24 (11), 4687–4702.

(11)

Petroutsos, D.; Tokutsu, R.; Maruyama, S.; Flori, S.; Greiner, A.; Magneschi, L.; Cusant, L.; Kottke, T.; Mittag, M.; Hegemann, P.; et al. A Blue-Light Photoreceptor Mediates the Feedback Regulation of Photosynthesis. Nature 2016, 537 (7621), 563–566.

(12)

Crosson, S.; Moffat, K. Structure of a Flavin-Binding Plant Photoreceptor Domain: Insights into Light-Mediated Signal Transduction. Proc. Natl. Acad. Sci. U. S. A. 2001, 98 (6), 2995–3000.

(13)

Fedorov, R.; Schlichting, I.; Hartmann, E.; Domratcheva, T.; Fuhrmann, M.; Hegemann, P. Crystal Structures and Molecular Mechanism of a Light-Induced Signaling Switch: The Phot-LOV1 Domain from Chlamydomonas Reinhardtii. Biophys. J. 2003, 84 (4), 2474–2482.

(14)

Möglich, A.; Ayers, R. A.; Moffat, K. Structure and Signaling Mechanism of Per-ARNT-Sim Domains. Structure 2009, 17 (10), 1282–1294. 20


Page 20 of 37

Page 21 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Photoreaction of Cr-Phototropin constructs (15)

Salomon, M.; Christie, J. M.; Knieb, E.; Lempert, U.; Briggs, W. R. Photochemical and Mutational Analysis of the FMN-Binding Domains of the Plant Blue Light Receptor, Phototropin. Biochemistry

2000, 39 (31), 9401–9410. (16)

Swartz, T. E.; Corchnoy, S. B.; Christie, J. M.; Lewis, J. W.; Szundi, I.; Briggs, W. R.; Bogomolni, R. A. The Photocycle of a Flavin-Binding Domain of the Blue Light Photoreceptor Phototropin. J. Biol. Chem. 2001, 276 (39), 36493–36500.

(17)

Conrad, K. S.; Manahan, C. C.; Crane, B. R. Photochemistry of Flavoprotein Light Sensors. Nat. Chem. Biol. 2014, 10 (10), 801–809.

(18)

Kasahara, M.; Swartz, T. E.; Olney, M. A.; Onodera, A.; Mochizuki, N.; Fukuzawa, H.; Asamizu, E.; Tabata, S.; Kanegae, H.; Takano, M.; et al. Photochemical Properties of the Flavin Mononucleotide-Binding Domains of the Phototropins from Arabidopsis, Rice, and Chlamydomonas Reinhardtii. Plant Physiol. 2002, 129 (2), 762–773.

(19)

Christie, J. M.; Swartz, T. E.; Bogomolni, R. A.; Briggs, W. R. Phototropin LOV Domains Exhibit Distinct Roles in Regulating Photoreceptor Function. Plant J. 2002, 32 (2), 205–219.

(20)

Tokutomi, S.; Matsuoka, D.; Zikihara, K. Molecular Structure and Regulation of Phototropin Kinase by Blue Light. Biochim. Biophys. Acta - Proteins Proteomics 2008, 1784 (1), 133–142.

(21)

Matsuoka, D.; Tokutomi, S. Blue Light-Regulated Molecular Switch of Ser/Thr Kinase in Phototropin. Proc. Natl. Acad. Sci. 2005, 102 (37), 13337–13342.

(22)

Harper, S. M.; Neil, L. C.; Gardner, K. H. Structural Basis of a Phototropin Light Switch. Science.

2003, 301 (5639), 1541–1544. (23)

Pfeifer, A.; Majerus, T.; Zikihara, K.; Matsuoka, D.; Tokutomi, S.; Heberle, J.; Kottke, T. Time-Resolved Fourier Transform Infrared Study on Photoadduct Formation and Secondary Structural Changes within the Phototropin LOV Domain. Biophys. J. 2009, 96 (4), 1462–1470.

(24)

Koyama, T.; Iwata, T.; Yamamoto, A.; Sato, Y.; Matsuoka, D.; Tokutomi, S.; Kandori, H. Different Role of the Jalpha Helix in the Light-Induced Activation of the LOV2 Domains in Various Phototropins. Biochemistry 2009, 48 (32), 7621–7628.

(25)

Oide, M.; Okajima, K.; Kashojiya, S.; Takayama, Y.; Oroguchi, T.; Hikima, T.; Yamamoto, M.; Nakasako, M. Blue Light-Excited Light-Oxygen-Voltage-Sensing Domain 2 (LOV2) Triggers a Rearrangement of the Kinase Domain to Induce Phosphorylation Activity in Arabidopsis Phototropin1. J. Biol. Chem. 2016, 291 (38), 19975–19984.

(26)

Eitoku, T.; Nakasone, Y.; Matsuoka, D.; Tokutomi, S.; Terazima, M. Conformational Dynamics of Phototropin 2 LOV2 Domain with the Linker upon Photoexcitation. J. Am. Chem. Soc. 2005, 127 (38), 13238–13244.

(27)

Nakasone, Y.; Eitoku, T.; Matsuoka, D.; Tokutomi, S.; Terazima, M. Dynamics of Conformational Changes of Arabidopsis Phototropin 1 LOV2 with the Linker Domain. J. Mol. Biol. 2007, 367 (2), 432–442.

(28)

Takeda, K.; Nakasone, Y.; Zikihara, K.; Tokutomi, S.; Terazima, M. Dynamics of the Amino-Terminal and Carboxyl-Terminal Helices of Arabidopsis Phototropin 1 LOV2 Studied by the Transient Grating. J. Phys. Chem. B 2013, 117 (49), 15606–15613. 21




Takakado, A.; Nakasone, Y.; Okajima, K.; Tokutomi, S.; Terazima, M. Light-Induced Conformational Changes of LOV2-Kinase and the Linker Region in Arabidopsis Phototropin2. J. Phys. Chem. B 2017, 121 (17), 4414–4421.

(30)

Halavaty, A. S.; Moffat, K. Coiled-Coil Dimerization of the LOV2 Domain of the Blue-Light Photoreceptor Phototropin 1 from Arabidopsis Thaliana. Acta Crystallogr. Sect. F. Struct. Biol. Cryst. Commun. 2013, 69, 1316–1321.

(31) Terazima, M. Diffusion Coefficients as a Monitor of Reaction Kinetics of Biological Molecules. Phys. Chem. Chem. Phys. 2006, 8 (5), 545–557. (32)

Terazima, M. Studies of Photo-Induced Protein Reactions by Spectrally Silent Reaction Dynamics Detection Methods: Applications to the Photoreaction of the LOV2 Domain of Phototropin from Arabidopsis Thaliana. Biochim. Biophys. Acta - Proteins Proteomics 2011, 1814 (8), 1093–1105.

(33)

Terazima, M. Time-Dependent Intermolecular Interaction during Protein Reactions. Phys. Chem. Chem. Phys. 2011, 13 (38), 16928.

(34)

Nakasako, M.; Zikihara, K.; Matsuoka, D.; Katsura, H.; Tokutomi, S. Structural Basis of the LOV1 Dimerization of Arabidopsis Phototropins 1 and 2. J. Mol. Biol. 2008, 381 (3), 718–733.

(35)

Nakasone, Y.; Kawaguchi, Y.; Kong, S.-G.; Wada, M.; Terazima, M. Photoinduced Oligomerization of Arabidopsis Thaliana Phototropin 2 LOV1. J. Phys. Chem. B 2014, 118 (49), 14314–14325.

(36)

Nakasone, Y.; Zikihara, K.; Tokutomi, S.; Terazima, M. Photochemistry of Arabidopsis Phototropin 1 LOV1: Transient Tetramerization. Photochem. Photobiol. Sci. 2013, 12 (7), 1171.

(37)

Pfeifer, A.; Mathes, T.; Lu, Y.; Hegemann, P.; Kottke, T. Blue Light Induces Global and Localized Conformational Changes in the Kinase Domain of Full-Length Phototropin. Biochemistry 2010, 49 (5), 1024–1032.

(38)

Okajima, K.; Aihara, Y.; Takayama, Y.; Nakajima, M.; Kashojiya, S.; Hikima, T.; Oroguchi, T.; Kobayashi, A.; Sekiguchi, Y.; Yamamoto, M.; et al. Light-Induced Conformational Changes of LOV1 (Light Oxygen Voltage-Sensing Domain 1) and LOV2 Relative to the Kinase Domain and Regulation of Kinase Activity in Chlamydomonas Phototropin. J. Biol. Chem. 2014, 289 (1), 413– 422.

(39)

Peter, E.; Dick, B.; Stambolic, I.; Baeurle, S. A. Exploring the Multiscale Signaling Behavior of phototropin1 from Chlamydomonas Reinhardtii Using a Full-Residue Space Kinetic Monte Carlo Molecular Dynamics Technique. Proteins Struct. Funct. Bioinforma. 2014, 82 (9), 2018–2040.

(40)

Onodera, A.; Kong, S.-G.; Doi, M.; Shimazaki, K. I.; Christie, J.; Mochizuki, N.; Nagatani, A. Phototropin from Chlamydomonas Reinhardtii Is Functional in Arabidopsis Thaliana. Plant Cell Physiol. 2005, 46 (2), 367–374.

(41)

Okajima, K. Molecular Mechanism of Phototropin Light Signaling. J. Plant Res. 2016, 129 (2), 149– 157.

(42)

Aihara, Y.; Yamamoto, T.; Okajima, K.; Yamamoto, K.; Suzuki, T.; Tokutomi, S.; Tanaka, K.; Nagatani, A. Mutations in N-Terminal Flanking Region of Blue Light-Sensing Light-Oxygen and Voltage 2 (LOV2) Domain Disrupt Its Repressive Activity on Kinase Domain in the Chlamydomonas Phototropin. J. Biol. Chem. 2012, 287 (13), 9901–9909. 22


Page 22 of 37

Page 23 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60



Kottke, T.; Heberle, J.; Hehn, D.; Dick, B.; Hegemann, P. Phot-LOV1: Photocycle of a Blue-Light Receptor Domain from the Green Alga Chlamydomonas Reinhardtii. Biophys. J. 2003, 84 (2), 1192– 1201.

(44)

Kottke, T.; Hegemann, P.; Dick, B.; Heberle, J. The Photochemistry of the Light-, Oxygen-, and Voltage-Sensitive Domains in the Algal Blue Light Receptor Phot. Biopolymers 2006, 82 (4), 373– 378.

(45)

Nakasone, Y.; Eitoku, T.; Matsuoka, D.; Tokutomi, S.; Terazima, M. Kinetic Measurement of Transient Dimerization and Dissociation Reactions of Arabidopsis Phototropin 1 LOV2 Domain. Biophys. J. 2006, 91 (2), 645–653.

(46)

Tyrrell, H. J. V. Diffusion in Liquids; Butterworth: London, 1984.

(47)

Cussler, E. L. Diffusion; Cambridge University Press: Cambridge, 1994.

(48)

Buttani, V.; Losi, A.; Eggert, T.; Krauss, U.; Jaeger, K. E.; Cao, Z.; Gärtner, W. Conformational Analysis of the Blue-Light Sensing Protein YtvA Reveals a Competitive Interface for LOV-LOV Dimerization and Interdomain Interactions. Photochem. Photobiol. Sci. 2007, 6 (1), 41–49.

(49)

Choi, S.; Nakasone, Y.; Hellingwerf, K. J.; Terazima, M. Photochemical Reactions of the LOV and LOV-Linker Domains of the Blue Light Sensor Protein YtvA. Biochemistry 2016, 55 (22), 3107– 3115.

(50)

Kutta, R. J.; Hofinger, E. S. A.; Preuss, H.; Bernhardt, G.; Dick, B. Blue-Light Induced Interaction of LOV Domains from Chlamydomonas Reinhardtii. Chembiochem 2008, 9 (12), 1931–1938.

(51)

Von Smoluchowski, M. Versuch Einer Mathematischen Theorie Der Koagulationskinetik. Phys Chem 1917, 92 (1912), 156.

(52)

Yamamoto, A.; Iwata, T.; Sato, Y.; Matsuoka, D.; Tokutomi, S.; Kandori, H. Light Signal Transduction Pathway from Flavin Chromophore to the Jα Helix of Arabidopsis Phototropin1. Biophys. J. 2009, 96 (7), 2771–2778.

(53)

Harper, S. M.; Christie, J. M.; Gardner, K. H. Disruption of the LOV−Jα Helix Interaction Activates Phototropin Kinase Activity. Biochemistry 2004, 43 (51), 16184–16192.

(54)

Takahashi, F. Blue-Light-Regulated Transcription Factor, Aureochrome, in Photosynthetic Stramenopiles. J. Plant Res. 2016, 129 (2), 189–197.

(55)

Takahashi, F.; Yamagata, D.; Ishikawa, M.; Fukamatsu, Y.; Ogura, Y.; Kasahara, M.; Kiyosue, T.; Kikuyama, M.; Wada, M.; Kataoka, H. AUREOCHROME, a Photoreceptor Required for Photomorphogenesis in Stramenopiles. Proc. Natl. Acad. Sci. 2007, 104 (49), 19625–19630.

(56)

Freddolino, P. L.; Gardner, K. H.; Schulten, K. Signaling Mechanisms of LOV Domains: New Insights from Molecular Dynamics Studies. Photochem. Photobiol. Sci. 2013, 12 (7), 1158.

(57)

Okajima, K.; Kashojiya, S.; Tokutomi, S. Photosensitivity of Kinase Activation by Blue Light Involves the Lifetime of a Cysteinyl-Flavin Adduct Intermediate, S390, in the Photoreaction Cycle of the LOV2 Domain in Phototropin, a Plant Blue Light Receptor. J. Biol. Chem. 2012, 287 (49), 40972–40981.

(58)

Dasgupta, A.; Chen, C. H.; Lee, C.; Gladfelter, A. S.; Dunlap, J. C.; Loros, J. J. Biological Significance of Photoreceptor Photocycle Length: VIVID Photocycle Governs the Dynamic 23



Photoreaction of Cr-Phototropin constructs VIVID-White Collar Complex Pool Mediating Photo-Adaptation and Response to Changes in Light Intensity. PLOS Genet. 2015, 11 (5), e1005215. (59)

Pudasaini, A.; Shim, J. S.; Song, Y. H.; Shi, H.; Kiba, T.; Somers, D. E.; Imaizumi, T.; Zoltowski, B. D. Kinetics of the LOV Domain of ZEITLUPE Determine Its Circadian Function in Arabidopsis. Elife 2017, 6, e21646.

(60)

Möglich, A.; Moffat, K. Structural Basis for Light-Dependent Signaling in the Dimeric LOV Domain of the Photosensor YtvA. J. Mol. Biol. 2007, 373 (1), 112–126.

(61)

Diensthuber, R. P.; Bommer, M.; Gleichmann, T.; Möglich, A. Full-Length Structure of a Sensor Histidine Kinase Pinpoints Coaxial Coiled Coils as Signal Transducers and Modulators. Structure

2013, 21 (7), 1127–1136. (62)

Circolone, F.; Granzin, J.; Jentzsch, K.; Drepper, T.; Jaeger, K. E.; Willbold, D.; Krauss, U.; Batra-Safferling, R. Structural Basis for the Slow Dark Recovery of a Full-Length LOV Protein from Pseudomonas Putida. J. Mol. Biol. 2012, 417 (4), 362–374.

(63)

Röllen, K.; Granzin, J.; Panwalkar, V.; Arinkin, V.; Rani, R.; Hartmann, R.; Krauss, U.; Jaeger, K.-E.; Willbold, D.; Batra-Safferling, R. Signaling States of a Short Blue-Light Photoreceptor Protein PpSB1-LOV Revealed from Crystal Structures and Solution NMR Spectroscopy. J. Mol. Biol. 2016, 428 (19), 3721–3736.

(64)

Zoltowski, B. D.; Schwerdtfeger, C.; Widom, J.; Loros, J. J.; Bilwes, A. M.; Dunlap, J. C.; Crane, B. R. Conformational Switching in the Fungal Light Sensor Vivid. Science. 2007, 316 (5827), 1054– 1057.

(65)

Vaidya, A. T.; Chen, C.-H.; Dunlap, J. C.; Loros, J. J.; Crane, B. R. Structure of a Light-Activated LOV Protein Dimer That Regulates Transcription. Sci. Signal. 2011, 4 (184), ra50–ra50.

(66)

Nakatani, Y.; Hisatomi, O. Molecular Mechanism of Photozipper, a Light-Regulated Dimerizing Module Consisting of the bZIP and LOV Domains of Aureochrome-1. Biochemistry 2015, 54 (21), 3302–3313.

(67)

Heintz, U.; Schlichting, I. Blue Light-Induced LOV Domain Dimerization Enhances the Affinity of Aureochrome 1a for Its Target DNA Sequence. Elife 2016, 5, e11860.

(68)

Endres, S.; Granzin, J.; Circolone, F.; Stadler, A.; Krauss, U.; Drepper, T.; Svensson, V.; Knieps-Grünhagen, E.; Wirtz, A.; Cousin, A.; et al. Structure and Function of a Short LOV Protein from the Marine Phototrophic Bacterium Dinoroseobacter Shibae. BMC Microbiol. 2015, 15 (1), 30.

(69)

Conrad, K. S.; Bilwes, A. M.; Crane, B. R. Light-Induced Subunit Dissociation by a Light–Oxygen– Voltage Domain Photoreceptor from Rhodobacter Sphaeroides. Biochemistry 2013, 52 (2), 378–391.

(70)

Jentzsch, K.; Wirtz, A.; Circolone, F.; Drepper, T.; Losi, A.; Gärtner, W.; Jaeger, K. E.; Krauss, U. Mutual Exchange of Kinetic Properties by Extended Mutagenesis in Two Short LOV Domain Proteins from Pseudomonas Putida. Biochemistry 2009, 48 (43), 10321–10333.

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Photoreaction of Cr-Phototropin constructs Table 1 Physical and reaction properties of Cr phot constructs determined by this study Oligomeric states of reactant and photo-product, time constants of the adduct formation (τadd), D-change (τD-change) and dark recovery (τrec), and diffusion coefficients of reactant, intermediate and product. For comparison, those parameters of At phot2 are also given in brackets 26,29. *1: Under highly concentrated conditions, Cr LOV1 forms a dimer in the dark and it dissociates into monomer upon photoexcitation. *2: At phot2 LOV1 and At phot2 LOV1-hinge are in equilibrium between monomer and dimer in the dark. ND: not determined.

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Photoreaction of Cr-Phototropin constructs FIGURE LEGENDS FIGURE 1. Schematic drawing of the domain architecture of Cr phot. (a) The five fragments used in this study are illustrated as bars. (b) Crystal structure of LOV1 domain of Cr phot (PDB ID: 1N9L). The FMN chromophore is shown in stick representation. (c) Absorption spectra of Cr LOV1 (blue) and LOV1-hinge (red) in the dark state (solid line) and the light state (broken line). (d) Absorption spectra of Cr LOV2 (blue), LOV2-linker (red) and hinge-LOV2 (green) in the dark state (solid line) and the light state (broken line). FIGURE 2. Observed TG signals upon photoexcitation of Cr LOV1 (a) TG signal of Cr LOV1 obtained at q2 = 1.7 × 1010 m−2. The black broken line shows the best fit to the data using Eq. 1 and Eq. 2. (b) q2 dependence on the diffusion signal of Cr LOV1. The signals are normalized in the initial part of the diffusion signal. The black broken lines show the best fits to the observed TG signals using Eq. 2. The signals are perfectly reproduced by the fitted curves. FIGURE 3. Observed TG signals of upon photoexcitation of Cr LOV1-hinge (a) TG signal of Cr LOV1-hinge obtained at q2 = 1.7 × 1010 m−2. Inset: An enlarged region of the TG signal observed over the time range of 100 ns–30 ms. The black line represents the best-fit curve obtained using Eq. 1 and Eq. 2. (b) q2 dependence on the diffusion signal of Cr LOV1-hinge. The black broken lines show the best fit to the observed TG signals using Eq. 2. FIGURE 4. Concentration dependence on the diffusion signal of Cr LOV1-hinge (a) Concentration dependence on the diffusion signal obtained at a small q2 condition (q2 = 1.7 × 1010 m−2). The concentrations are 400 (red), 320 (yellow), 240 (green), 160 (cyan) and 80 µM (blue). The signals were normalized by the signal intensity before the diffusion signal. (b) Concentration dependence on the diffusion signal obtained at a large q2 condition (q2 = 4.3 × 1012 m−2). The concentrations and colors are the same as those of (a). The signals were normalized by the signal intensity before the diffusion signal. The black broken lines represent the best-fit curves to the observed TG signals using Eq. 2. (c) The rate constants of D-change are plotted against the concentration. The fitted linear line is shown in red. FIGURE 5. CD spectra and light induced changes of LOV1 constructs (a) CD spectra of Cr LOV1 (blue) and Cr LOV1-hinge (red) in the dark (solid lines) and light (broken lines) states. The optical path length was 1.0 cm and the protein concentration for both samples was 0.5 µM. (b) Dark recovery of the CD intensity at 220 nm after blocking of blue light illumination (blue: LOV1; red: LOV1-hinge). The black broken lines represent the fitted curves obtained from the single exponential function. FIGURE 6. Observed TG signals of upon photoexcitation of LOV2 constructs (a) TG signals of Cr LOV2 (blue) and LOV2-linker (red) obtained at q2 = 1.3 × 1010 m−2. The black broken lines represent the best-fit curves to the observed TG signals for LOV2 (Eq. 1 and 4) and LOV2-linker (Eq. 1 and 2). (b) q2 dependence on diffusion signal of Cr LOV2-linker. The signals are normalized by the signal intensity before 26


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Photoreaction of Cr-Phototropin constructs the diffusion signal. The black lines represent the best-fit curves to the observed TG signals using Eq. 2. FIGURE 7. CD spectra and light induced changes of LOV2 constructs (a) CD spectra of Cr LOV2 (blue), LOV2-linker (red) and hinge-LOV2 (green) in the dark (solid lines) and light (broken lines) states. The optical path length was 1.0 cm and the protein concentration for all samples was 0.5 µM. (b) Dark recovery of the CD intensity at 220 nm after blocking of blue light illumination (blue: LOV2; red: LOV1-linker; green: hinge-LOV2). The black broken lines represent the fitted curves obtained from the double exponential function. FIGURE 8. Proposed reaction schemes of Cr LOV1 constructs (a) Cr LOV1 and (b) Cr LOV1-hinge. FIGURE 9. Proposed reaction schemes Cr LOV2 constructs (a) Cr LOV2, (b) Cr LOV2-linker and (c) Cr hinge-LOV2.

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Fig. 1

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Fig. 2

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Fig. 3

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Fig. 4

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Fig. 5

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Fig. 6

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Fig. 7

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Fig. 8

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Fig. 9

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Photoreaction of Cr-Phototropin constructs TOC Graphic

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Photoreaction Dynamics of LOV1 and LOV2 of Phototropin from

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