Light-Induced Conformational Changes of LOV2-Kinase and the

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Light-Induced Conformational Changes of the LOV2Kinase and the Linker Region in Arabidopsis Phototropin2 Akira Takakado, Yusuke Nakasone, Koji Okajima, Satoru Tokutomi, and Masahide Terazima J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.7b01552 • Publication Date (Web): 07 Apr 2017 Downloaded from http://pubs.acs.org on April 8, 2017

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The Journal of Physical Chemistry

Light-Induced Conformational Changes of the LOV2-Kinase and the Linker Region in Arabidopsis Phototropin2

Akira Takakado,† Yusuke Nakasone,† Koji Okajima,§ Satoru Tokutomi,§ and Masahide Terazima*,† †

Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan

§

Department of Biological Science, Graduate School of Science, Osaka Prefecture University, Sakai,

Osaka 599-8531, Japan

* Corresponding author Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan [email protected]

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ABSTRACT Phototropins are blue light sensors found in a variety of higher plants and algae. The photochemical reactions of this family of proteins have attracted much attention since their discovery. Phototropins have two light sensor domains called light-oxygen-voltage 1 (LOV1) and LOV2. After the formation of the characteristic adduct of the LOV domain, a conformational change of the C-terminal region of the LOV2 domain occurs, and characterizing this change is important for understanding biological function, i.e., kinase activation. Here, the reaction dynamics of the Jα-helix and the extended region adjacent to the Jα-helix (connector) have been investigated. The conformation of the connector part and the Jα-helix were found to alter significantly in two-state manner. Furthermore, the conformational change of the kinase domain was also successfully detected as a change in translational diffusion, although the CD intensity owing to the kinase domain movement was almost silent. These observations indicate that the tertiary structure of the kinase domain changes. The rate of the kinase domain change is almost the same as that of the change for the LOV2-linker, suggesting that the conformational change of the linker is the rate-determining step for kinase activation.


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INTRODUCTION Light is an essential environmental stimulus for animals, microorganisms and plants. Consequently, these organisms have developed several light sensor proteins. Phototropins (phots) are blue light sensors found widely in a variety of organisms. For example, in higher plants, phots regulate phototropism, stomatal opening and chloroplast relocation to ensure optimal photosynthesis.1–5 Most plants possess two isoforms of phot, designated as phot1 and phot2.6 Phot is a member of subfamily VIIIb of the AGC kinases (cAMP-dependent protein kinase, cGMP-dependent protein kinase and phospholipid-dependent protein kinase C).6,7 The phots consist of a kinase domain in the C-terminal half, and two light sensor domains called light-oxygen-voltage 1 (LOV1) and LOV2 located in the N-terminal half (Figure 1).8 The LOV domain contains a flavin mononucleotide (FMN), which functions as the non-covalently bound chromophore.8,9 Upon photoexcitation of the chromophore, the LOV domain undergoes a cysteinyl adduct formation between the FMN and a nearby cysteine residue.10,11 The adduct formation takes place within a few microseconds10 and recovers back to the dark state thermally over a time range of seconds to minutes.12 The photochemistry of the chromophore is well conserved among all LOV domains and is considered vital for signal transduction, although a recent report has suggested that the Cys-lacked VVD protein, a kind of LOV proteins, still undergoes signal transduction.13 The LOV2 domain is connected to the kinase domain via a linker region and this region is believed to regulate kinase activity.4,14 To understand the light-induced activation of the kinase domain, conformational changes are considered to be a key step; thus, the reactions after adduct formation of the LOV2 domain have been studied extensively. NMR studies revealed an interaction between the amphipathic helix known as the Jα-helix located within the C-terminal extended region (Fig. 1) and the LOV2 domain in the dark state, and it was shown that this interaction is disrupted upon blue light illumination for Avena sativa (As.) phot1 LOV2 domain with a 40-amino acid C-terminal extension (L404–H560).14 FT-IR experiment reported characteristic differences of the light-minus-dark-difference infrared 3 ACS Paragon Plus Environment


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spectra between Arabidopsis thaliana (At.) phot1 LOV2-core (K462–R586) and LOV2-Jα (K462– D617), indicating a conformational change (unfolding) within the Jα-helix by light illumination.15 Kinetics of the unfolding of the Jα-helix was studied for As. LOV2 with the Jα-helix (L404–H560) by time-resolved vibrational spectroscopy and the time constant of the full unfolding phase was 240 µs.16 Most of these studies focused on the conformational change of the Jα-helix. Conformation changes of the more extended region, linker region, were studied by the time-resolved transient grating (TG) technique, which elucidated the reaction dynamics of At. phots for LOV2-linker fragments (D363–H575) containing the LOV2 domain and the full linker region. The photoexcitation of phot2LOV2-linker leads to the linker domain dissociating from the LOV2 domain with a time constant of 140 µs and the linker helix unfolds with a time constant of 2 ms.17,18 The dissociation and subsequent unfolding are conserved in the phot1LOV2-linker with slight differences in the time constants (300 µs and 1 ms for dissociation and unfolding, respectively).19 In addition to changes of the linker helix, the importance of the A′α-helix, which locates to the N-terminal side of the LOV2 domain, was shown by biochemical and biophysical studies.20–22 Since the kinase is constitutively active in isolation,23 it is believed that the kinase domain is activated by a conformational change of the A′α-helix as well as the linker helix.24 In this research, we aimed to reveal two points on the reaction kinetics after the photoreaction of the At.phot2 LOV2 domain to understand the conformational change of the kinase domain. First, although the reaction kinetics of the phot2LOV2-linker (D363–L574) have been reported, the kinetics of the Jα-helix (E505–L529) and relationship between the Jα-helix and the linker outside of the Jα-helix (P530–F577; referred here as connector, Fig. 1) remain unresolved. Does the conformation of the connector part change beside that of the Jα-helix? If it does, how fast is the Jα-helix unfolding process? Is it the same as the conformational change of the linker? These points are important, because an essential role of the connector part has been recently reported by a phosphorylation assay for At. phot1.25 It was shown that point mutations located in the middle of the connector region (e.g., H629A, S630A and K636A of phot1LOV2-linker) reduce kinase activity 4 ACS Paragon Plus Environment

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in the light-irradiated condition. This result suggests the connector part and the Jα-helix are important in controlling kinase activity. However, there has been no report on the conformational change and the reaction dynamics of this region. To clarify this point, we measured the reaction kinetics of LOV2-Jα (D363–E528) consisting of the LOV2 domain and the Jα-helix and compared the results with those of the LOV2-linker (D363–L574). Second, while most previous studies on At.phots have focused on the adjacent helices of the LOV2 domain, the kinetics of kinase activation remain unclear. One of the reasons for the paucity of experimental data is the difficulty of producing and purifying a LOV2-kinase sample. However, recently, such a purification approach became available and a SAXS study reported that phot constructs containing the LOV2 domain and the kinase domain show a slight elongation of the overall structure upon light illumination.24,26–28 In this study, we investigated the kinetics of kinase activation by comparing several fragments of At. phot2 LOV2-kinase (D363–F915) containing the kinase domain for the first time.

EXPERIMENTAL SECTION Preparation of sample proteins. DNA fragments encoding Arabidopsis thaliana phototropin2 LOV2-Jα (D363–E528), LOV2-linker (D363–L574) and LOV2-kinase (D363–F915) were cloned into a pET-28a vector. For the LOV2-kinase sample, an Asp720Asn mutation in the middle of the kinase domain was introduced to eliminate its binding affinity for Mg-ATP.26 Target proteins were overexpressed in Escherichia coli. (E. coli) Rosetta2(DE3) cells (Novagen), which had been transformed with each expression vector. Harvested cells were re-suspended in HEPES buffer (20 mM HEPES, 0.5 M NaCl and 10 % (w/v) glycerol, pH 7.5) containing 1 mg/mL lysozyme (Sigma Aldrich) and a protease inhibitor cocktail (Nacalai Tesque) and disrupted by super-sonication. After centrifugation, the supernatant was loaded onto a His-Trap HP column (GE Healthcare), and the target protein was isolated by an imidazole gradient (30 to 500 mM) in HEPES buffer. The eluted samples were further purified by size exclusion chromatography (SEC) 5 ACS Paragon Plus Environment


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(Sepharose S-100HR, GE Healthcare). Tris buffer (20 mM Tris and 10 % (w/v) glycerol, pH 7.8) containing 100 mM NaCl was used for SEC. Although the LOV2-linker sample was sufficiently pure following SEC, the LOV2-Jα and the LOV2-kinase samples were not. Therefore ion-exchange chromatography (Mono Q, GE Healthcare) was used for further purification of these two samples. For ion-exchange chromatography, the Tris buffer without NaCl was used as the sample loading buffer, and a Tris buffer containing 1000 mM NaCl was used as the elution buffer. Finally, all samples were dissolved in Tris buffer containing 500 mM NaCl. All experiments were performed in this buffer. The concentration of the samples was determined by absorbance at 447 nm using the extinction coefficient of 13,500 M–1cm–1.12

Circular dichroism measurement. The secondary structures of the samples were examined by circular dichroism (CD) spectroscopy (J-720W1 JASCO). A quartz-cell (optical path length = 0.2 (for spectrum measurement) and 1 cm (for time-resolved measurement)) was used and the protein concentration was 4 µM (for spectrum measurement) and 1 µM (for time-resolved measurement) for all samples. To measure spectra of the light-adapted state of the protein, the samples were illuminated by a blue light-emitting diode (465 nm) during measurement. During the measurements, samples were stirred with a magnetic stirrer at 23 °C.

Transient grating and transient lens measurement. The experimental setup for the transient grating (TG) and the transient lens (TrL) experiments were similar to previous reports.17– 19,29

The wavelength of the pump beam was 462 nm and the probe beam was 835 nm (for TG

experiments) and 633 nm (for TrL experiment). A laser pulse from a dye laser (Lumomics, HyperDye 300) pumped by an eximer laser (Lambda Physik, XeCl operation) was used as a pump pulse. As a probe beam, a diode laser (CrystaLaser) was used for the TG measurements and a He-Ne laser was used for the TrL measurement. A quartz-cell (optical path length = 2 mm) was used. The repetition rate of the pump pulse was 0.03 Hz. This repetition rate is sufficiently slow for 6 ACS Paragon Plus Environment

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completion of the dark recovery of the LOV2 domain (6 s). All measurements were performed at 23 °C.

Principles of TG and TrL. The principle of the TG method TrL methods have been described in detail previously.30–32 Two excitation beams induce a temporal interference pattern characterized by a grating wave number q, which is determined by the wavelength of the excitation laser and the crossing angle of two beams. The TG signal is generated by a photo-induced refractive index change (δn) created by this light. The signal consists of the thermal grating (δnth) because of a change in temperature due to released thermal energy, and the species grating arises from different chemical species. The species grating component consists of the reactant component (δnR) and the product component (δnP). Consequently, the TG signal (ITG(t)) is expressed by:

= { − + }

(1)

where α is a constant. Here, the product indicates chemical species created by light illumination, including transient intermediates. The thermal grating decays by the thermal diffusion process at a rate constant of Dthq2, where Dth represents a thermal diffusivity of the solution and q is the grating wavenumber. The time-dependence of the species grating comes from the molecular diffusion and chemical reaction. When the chemical reaction is ignorable during the observing time-scale, ITG(t) is expressed by:

= { − − exp − + exp − }

(2)

where , and are the initial refractive index changes due to the thermal grating, reactant and the product, respectively. DR and DP represent the diffusion coefficients of the reactant and product, respectively. 7 ACS Paragon Plus Environment


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When the reaction takes place in the observation time window, this kinetic is included in the temporal profile of the signal. For example, when the reaction takes following scheme with a rate constant (k),

$

!"""""# !""""# %

(Scheme 1)

where R, I and P are the reactant, the intermediate and the final product, respectively, the molecular diffusion signal is written by:

= &− − + ' ( − * )

*+ ,*-

)

1 {−( / + ,*- . 0 )

− 3 . /0)

+ 2 } + (3)

where ( and DI are the initial refractive index change and the diffusion coefficient of the intermediate species, respectively. By a q2 dependence of the TG signal, one can determine the rate constant k and the other parameters. In the TrL method, the signal arises from the refractive index change created by the (nearly) Gaussian beam. The origins of the refractive index change are similar to the TG method. The TrL signal consists of the thermal effect (thermal lens) and a change to the chemical species (species lens).

RESULTS AND DISCUSSION Conformational changes of the Jα and linker regions (a) Diffusion change. The reaction dynamics of LOV2-Jα were studied by the TG method. A typical TG signal of LOV2-Jα is shown in Fig. 2(a). The qualitative features are essentially similar to that of LOV2-linker and the assignment of the signal components were reported 8 ACS Paragon Plus Environment

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previously.17,19 Briefly, the signal rises quickly within the response time of this system, then the signal shows a decay-rise profile within a few microseconds and decays within 100 microseconds. This time dependence was expressed well by a biexponential function with a q-independent time constants of 0.9 µs, and a q-dependent time constant (Dthq2)-1 (=9.9 µs). Since only the diffusional process depends on the grating wavenumber q, this q dependence indicates that these phases are attributed to the adduct formation and the decay of the thermal grating (δnth) component, respectively.10,17 The time range of the next rise-decay component (100 µs to 1 s in Fig. 2(a)) was dependent on q2 (Fig. 2(b)), indicating that this component represents a protein diffusion process (diffusion signal). Comparing the signs of the refractive index change of the thermal grating (δnth < 0) and of the diffusion signal enabled us to determine the signs of the refractive index changes of the rise and decay components as negative and positive, respectively. From these signs and eq. 2, the rise and decay components of the diffusion signal were assigned to the diffusions of the reactant and the product, respectively. Because the rate of the rise component is faster than that of the decay component, D of the product is apparently smaller than that of the reactant; i.e., DR > DP. We first determined DR and DP from a signal taken over a sufficiently long time range (20 ms to 1 s). Because the reaction rate was determined to be 500 µs as described below, the kinetics can be neglected in this time region. We fitted the signal using the biexponential function (eq. 2) to yield values of DR and DP of 9.4 × 10–11 m2/s and 8.4 × 10–11 m2/s, respectively. Previous studies on LOV2 and LOV2-linker showed that D of LOV2-linker decreases from DR = 8.8 × 10–11 m2/s to DP = 6.8 × 10–11 m2/s upon formation of the light state, whereas D of LOV2 slightly increases from DR = 10.3 × 10–11 m2/s to DP = 10.5 × 10–11 m2/s.17,18 DR of LOV2-Jα is larger than that of the LOV2-linker but smaller than that of LOV2. This order of D is the same as the order of molecular size and it is reasonable according to the Stokes-Einstein relationship. To investigate the origin of the D change to LOV2-Jα, we have used CD spectroscopy to probe for secondary structure changes. Figure 3(a) shows CD spectra of the LOV2-Jα and the LOV2-linker under dark and light conditions. Both samples show a decrease in the intensity of CD 9 ACS Paragon Plus Environment


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spectra upon light illumination. These changes are mainly attributed to the disruption of the helical structure, which has a characteristic peak at 208 and 222 nm.29 The light-induced change of the CD intensity at 222 nm was monitored by the time-dependent change after blocking light illumination (Fig. 3(b)). Both signals were well reproduced by a single exponential function and the lifetimes of the light states were estimated to be 6 ± 1 s. This lifetime agrees well with that of the recovery process of the chromophore (6 s) detected by the change in absorption intensity at 447 nm, indicating that the secondary structures are restored concomitantly with the recovery of the chromophore. The amplitudes of signals depicted in Fig. 3(b), which represent the amount of secondary structure change by light illumination, were 0.85 and 3.8 mdeg for LOV2-Jα and LOV2-linker, respectively. Thus, the LOV2-Jα exhibits a much smaller change than the LOV2-linker. The reduction of D for the LOV2-linker has been attributed to the unfolding of the linker helix.17,19,29 Similarly for LOV2-Jα, the decrease of D was attributed to the unfolding of the secondary structure of the Jα-helix, which was confirmed by the CD measurement. The reduction of D implies an increase in friction during the diffusion process. The enhanced friction (∆f) of LOV2-Jα is calculated to be 5.2 × 10–12 kg/s based on the equation ∆f = kBT{1/DP – 1/DR}, where kB is the Boltzmann constant and T is the temperature. It is interesting to note that the enhanced friction of LOV2-Jα is much smaller than that of LOV2-linker (∆f = 1.4 × 10–11 kg/s). If we assume that this increase in the friction comes from the unfolding of the helices, we can estimate roughly the extent of unfolding of the linker region as follows. The number of amino acid residues of the Jα-helix is 25. Since the average increase in friction per amino acid reside is 2.1 × 10–13 kg/s, the number of the unfolded helix in the linker should be 1.4 × 10–11/2.1 × 10–13 = 66. Unfortunately, the secondary structure of the connector region is not known. Using the Phyre2 program33 to predict the secondary structure of the P530–H575 region, we obtain that sequences P536–H543 and S558– A567 may form helical structures, which would give a helix length of 18+25=43 residues in the linker region. This number is smaller than the estimation of 66 residues based on changes in friction. 10 ACS Paragon Plus Environment

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We consider that most of the connector region possesses a helical structure due to interaction with the LOV domain in the dark state, but is intrinsically disordered in the light state, which is similar to the Jα-helix case. Large changes in D and the CD spectrum for the LOV2-linker should indicate that the helix structure within the connector region in addition to the Jα-helix becomes disordered upon light illumination. As shown in the introduction section, the importance of the connector region for kinase activity has been revealed. The conformational change of the connector region detected in this study is likely to regulate kinase activity.

(b) Kinetics. To determine the reaction kinetics of LOV2-Jα, we measured the diffusion signals at several q2 values (Figure 2(b)). The TG signals were normalized by the number of excited molecules, which was determined by the intensity of the species grating after thermal diffusion. The diffusion signal intensity was significantly dependent on q2; i.e., the signal was weak in a fast time range (large q2) and became stronger as the observation time range increased. This q2 dependence (observation time range dependence) indicates that D of the product is gradually changing in the observation time range. The observed TG signal can be reproduced very well over a wide observation time range (i.e., at various q2) using eq. 3 with two adjustable parameters, the diffusion coefficient of the intermediate DI and the reaction rate, k. DI was determined to be 9.4 × 10–11 m2/s, and the time constant of the D change was determined to be 500 µs. This time constant of the D change of the LOV2-Jα (500 µs) is faster than that of the LOV2-linker (2 ms).17 If the disordering of connector follows sequentially the conformational change of the Jα-helix, the conformational change of the linker region should be expressed by a two-step process (i.e., with two time constants). However, the TG signal of the LOV2-linker17,19,29 and LOV2-Jα were reproduced well by one time constant. Hence, the conformations of the Jα-helix and connector part should change simultaneously. There may be two possible explanations for the slow conformational change of the linker compared with that of the Jα-helix. Firstly, although unfolding 11 ACS Paragon Plus Environment


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of the Jα-helix regulates the conformation change of the connector, the rate is slower for the LOV2-linker due to the large and massive connector part. Secondly, both the Jα and connector interact with the LOV2 domain, and the rate is slower due to the stronger domain interaction between LOV2 and the linker than between LOV2 and the Jα-helix. We cannot clearly determine which explanation is correct. On the basis of the crystal structure of At. phot1 LOV2, however, a possible interaction between the connector and the LOV2 was proposed.34 Hence, the later explanation may be plausible.

Conformational change of the kinase domain. Figure 4 shows typical TG signals of LOV2-kinase and the LOV2-linker. The profile of the TG signal of LOV2-kinase is very similar to that of the LOV2-linker. The initial decay-rise-decay components (10 ms), which is the molecular diffusion signal, shifts to a slower time and the peak intensity increases in the presence of the kinase domain. It indicates that the diffusion coefficients of LOV2-kinase are different from those of LOV2-linker. Figure 5 shows the q2 dependence of the diffusion signal of LOV2-kinase. From fitting the signal obtained at small q2 the DR and DP values were determined to be 7.3 × 10–11 m2/s and 3.9 × 10–11 m2/s, respectively. The DR of LOV2-kinase is smaller than that of the LOV2-linker (DR = 8.8 × 10–11 m2/s), which is reasonable, because the molecular size of the LOV2-kinase is larger. A significant large D change was observed for LOV2-kinase (DR = 7.3 × 10–11 m2/s and DP = 3.9 × 10–11 m2/s) when compared with that of the previously reported values for the LOV2-linker (DR = 8.8 × 10–11 m2/s and DP = 6.8 × 10–11 m2/s).17 From the same calculation for LOV2-Jα, the increase in friction was determined as –11

∆f = 5.4 × 10

kg/s. A significant change in ∆f for LOV2-kinase compared to that of LOV2-linker

(∆f = 1.4 × 10–11 kg/s) indicates the larger conformational change of LOV2-kinase, suggesting that 12 ACS Paragon Plus Environment

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the kinase domain undergoes significant conformational change. Interestingly, however, this larger change was not detected by the CD measurement. The CD spectrum of LOV2-kinase is shown in Figure 3(a). Similar to the LOV2-linker case, the CD intensity at 222 nm of LOV2-kinase decreased upon light illumination. Amplitudes of signals depicted in Figure 3(b), which represent the amount of the secondary structural change, were 3.8 mdeg and 4.0 mdeg for the LOV2-linker and the LOV2-kinase, respectively. The similar decrease in the CD intensity for the LOV2-linker and LOV2-kinase suggests that the secondary structure change of LOV2-kinase is limited to the linker helices region; i.e., the secondary structure in the kinase domain does not change significantly. The fact that the light-induced change in the CD intensity of LOV2-kinase was similar to that of the LOV2-linker seems to be in contradiction with the large D-change. What causes the decrease in D? We consider that the D change may be attributed to a tertiary structure change, which may enhance the interaction with solvent and/or to a change in molecular shape leading to a decrease of D. We consider that the D measurement will be a useful tool to detect conformational changes of kinase domains. The kinase domain consists of two lobes: a smaller N-terminal lobe containing an ATP-binding site and a larger C-terminal lobe containing the substrate-binding site.6 One clear structural difference between PKA, which is a typical AGC VIIIb protein kinase, and the kinase domain of phot is the presence of the additional loop structure called the activation loop (A-loop) in the C-terminal lobe of LOV2-kinase.6 An FT-IR study35 and a molecular simulation study36 have proposed that the conformational change (unfolding35/stretching36) of the A-loop takes place during kinase activation, and hence, the A-loop has been postulated to play an important role for biological function. A previous FT-IR study on full-length phot from Chlamydomonas reinhardtii suggested that the helical structure in the A-loop extension is disrupted by light illumination.35 Considering these previous studies, the conformation change observed here may reflect a conformational change of the A-loop extension within the kinase domain. To examine the possibility of the structural change in the A-loop extension, we tried to measure photoreactions of a mutant, in which the 13 ACS Paragon Plus Environment


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A-loop extension (S724–P757) is substituted to four glycine residues. The TG signal of the mutated LOV2-kinase at q2 = 7.8 × 1011 m–2 is shown in Fig. S1 in Supporting Information. We fitted the signal using the biexponential function (eq. 2) and determined DR and DP to be 6.8 × 10–11 m2/s and DP = 3.2 × 10–11 m2/s, respectively. The DR is similar to that of the wild-type and the D change resembles that of the LOV2-kinase when compared with the change of the LOV2-linker. This result indicates that the conformational change of the A-loop region is minor. We tentatively consider that the main contribution of the D change of the LOV2-kinase may be relocation of the kinase domains against the LOV2-linker part. A SAXS study revealed that the LOV2-kinase forms an elongated conformation upon light illumination (The radius of gyration was 32.4 Å (dark) and 34.8 Å (light)).26 The slightly elongated conformation may represent the changes in the linker region as well as the movement of the kinase domain. Exposure of amino acid residues in the kinase domain upon light illumination is another possible explanation for the D change. A docking study proposed that the LOV2 domain may dock onto the catalytic cleft within the kinase domain and this docking inhibits kinase activity.6 The kinase activity may be regulated by the docking/undocking of the LOV2 domain from the catalytic cleft. A crystal structure for a homologue PKA showed that it possesses closed and opened conformations.37 These conformations may exist for this phototropin kinase and such open-close motion of the kinase domain may cause the D reduction observed here. To determine the reaction kinetics of LOV2-kinase, we measured TG signals at several q2 values (Fig. 5) and analyzed the TG signal by a similar method described in the preceding section. The time development of the diffusion signal was well reproduced by eq. 3. The diffusion coefficient of the intermediate (DI) and time constant of the D decrease were determined from the q2 dependence of the diffusion signal to be 7.3 × 10–11 m2/s and 2 ms, respectively. It is rather surprising that the reaction rate of unfolding of the linker helix is not affected by the presence/absence of the kinase domain. The same rate constant of LOV2-kinase to that of the LOV2-linker suggests that the origin of the D-change for LOV2-kinase is caused by the 14 ACS Paragon Plus Environment

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conformational change in the linker region. This result suggests that the former explanation (relocation of the kinase domains against the LOV2-linker part) is more plausible for the origin of the D-change. The influence of the kinase domain on the reaction phase was investigated by the transient lens (TrL) technique. In principle, we can monitor the volume change process as a change of refractive index by the TG experiment. However, the strong molecular diffusion signal may mask possible volume changes, if rates of the volume changes were overlapped with the diffusion signal. To overcome this difficulty of detection, the TrL technique was applied. Since the time range of the molecular diffusion process in the TrL signal is much slower than that of the TG signal, the volume change may be detected clearly. Indeed, the TrL measurements have previously revealed that the linker regions undergo conformational changes with time constants of 140 µs (phot2LOV2-linker)18 and 300 µs (phot1LOV2-linker),19 which are attributed to volume changes associated with the dissociations of the linker domains from the LOV2 cores. The TrL signal of the LOV2-kinase sample is shown in Fig. S2. Upon photo-excitation, the TrL signal showed the thermal lens component, which decays with a time constant of ~100 ms (almost constant in Fig. S2) and another phase in the faster time region (> 1 ms). Since the fast decay is neither the thermal lens component nor the population lens due to the absorption change, we attribute the decay signal to the kinetics of the volume lens components. The signal was reproduced by a single exponential function beside the thermal lens component and the time constant was determined to be 330 µs. The time constant is similar to the previously reported time constant for LOV2-linker samples. The relatively slower time constant may be due to the presence of the kinase domain, that is, the mobility of linker region is slightly suppressed by the presence of the kinase domain. Schematic illustration of the conformational change of the LOV2-kinase is depicted in Figure 6. After the photochemistry of chromophore (0.9 µs), the Jα-helix dissociates from the LOV2 domain (330 µs). This may destabilize the helical structure of the linker, leading to their unfolding (2 ms). The conformational change of the kinase is concomitantly induced, which should be 15 ACS Paragon Plus Environment


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relevant for its activation. In summary, we found that the LOV2-linker exhibits much larger changes in both the D-value and CD intensity when compared with those of LOV2-Jα, indicating that the conformation of the connector part changes significantly, more than that of the Jα-helix, upon light irradiation. This movement should regulate the kinase activity. The conformation of the connector and Jα-helix change simultaneously, but the rate is slower for the LOV2-linker (2 ms) than that of LOV2-Jα (500 µs). Recently, phototropin LOV-Jα was used as an optogenetical tool by utilizing the conformational change of the LOV-Jα.38,39 The LOV2-linker fragment containing the connector part may be useful for robust regulation in optogenetics. We also successfully detected the conformational change dynamics of the kinase domain and determined the rate for the first time. The conformational change of the kinase occurs simultaneously with that of the linker, indicating that the unfolding of the linker helix is a rate-determining step of kinase activation.

ACKNOWLEDGMENTS 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 from the Ministry of Education, Culture, Sports, Science and Technology in Japan (to M.T.) and in part by Grants-in-aid for Scientific Research on Innovative Areas 22120005 (to S. T.).


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Figure captions

Figure 1. A schematic drawing of the domain configuration of At. phot2. The three fragments used in this study (LOV2-Jα, LOV2-linker and LOV2-kinase) are illustrated as bars. The orange region is the connector region.

Figure 2. (a) A typical TG signal of LOV2-Jα at a grating number of q2 = 7.2 × 1011 m–2 and a concentration of 80 µM. (b) The q2 dependence of the molecular diffusion signals of LOV2-Jα at a concentration of 35 µM. The grating wavenumbers are q2 = 9.8 × 1012 m–2 (purple), 2.4 × 1012 m–2 (blue), 1.1 × 1012 m–2 (green) and 3.9 × 1011 m–2 (red). The fitted curves based on Scheme 1 (eq. 3) are shown as black lines.

Figure 3. (a) CD spectra of LOV2-kinase (blue), LOV2-linker (red) and LOV2-Jα (green) in the dark (solid line) and in the presence of light (dashed line). The protein concentration was 1 µM for all samples. (b) Temporal profiles of the CD intensities at 222 nm after terminating the light illumination for LOV2-kinase (blue), LOV2-linker (red) and LOV2-Jα (green). Each signal is fitted by a single exponential function (black line). These profiles represent the dark recovery dynamics of the secondary structure.

Figure 4. Typical TG signals of LOV2-kinase (blue) and LOV2-linker (red) at q2 = 3.3 × 1010 m–2 and a concentration of 20 µM. The inset shows the early time range of the signals.

Figure 5. The q2 dependence of the molecular diffusion signals of LOV2-kinase at a concentration of 100 µM. The grating wavenumbers are q2 = 1.1 × 1013 m–2 (purple), q2 = 3.0 × 1012 m–2 (blue), q2 = 9.5 × 1011 m–2 (green) and q2 = 2.9 × 1011 m–2 (red). The fitted curves using the two-state model (eq. 3) are shown as black lines. 22 ACS Paragon Plus Environment

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Figure 6. Schematic illustration of the conformational changes of LOV2-kinase upon blue light illumination.



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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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Table of Contents


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Light-Induced Conformational Changes of LOV2-Kinase and the

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