Dynamics of the Amino-Terminal and Carboxyl-Terminal Helices of

Aug 9, 2013 - Recently, conformational changes of the amino-terminal helix (A′α helix), .... of an Initial Step of Phototropin Signaling in Stomata...
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Dynamics of the Amino-Terminal and Carboxyl-Terminal Helices of Arabidopsis Phototropin 1 LOV2 Studied by the Transient Grating Kimitoshi Takeda,† Yusuke Nakasone,† Kazunori Zikihara,‡ Satoru Tokutomi,‡ and Masahide Terazima*,† †

Department of Chemistry, Graduate School of Science, Kyoto University, Kitashirakawa, Kyoto 606-8502, Japan Research Institute for Advanced Science and Technology, Department of Biological Science, Graduate School of Science, Osaka Prefecture University, Sakai, Osaka 599-8531, Japan

‡

ABSTRACT: Recently, conformational changes of the amino-terminal helix (A′α helix), in addition to the reported conformational changes of the carboxyl-terminal helix (Jα helix), have been proposed to be important for the regulatory function of the light-oxygen-voltage 2 domain (LOV2) of phototropin 1 from Arabidopsis. However, the reaction dynamics of the A′α helix have not been examined. Here, the unfolding reactions of the A′α and Jα helices of the LOV2 domain of phototropin 1 from Arabidopsis thaliana were investigated by the time-resolved transient grating (TG) method. A mutant (T469I mutant) that renders the A′α helix unfolded in the dark state showed unfolding of the Jα helix with a time constant of 1 ms, which is very similar to the time constant reported for the wildtype LOV2-linker sample. Furthermore, a mutant (I608E mutant) that renders the Jα helix unfolded in the dark state exhibited an unfolding process of the A′α helix with a time constant of 12 ms. On the basis of these experimental results, it is suggested that the unfolding reactions of these helices occurs independently.

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INTRODUCTION Phototropins (phot1 and phot2 in Arabidopsis thaliana) are blue-light sensor proteins that regulate phototropism, chloroplast movement, and stomata opening of higher plants.1−7 This protein consists of a kinase domain at the C-terminus, two light-oxygen-voltage (LOV) (LOV1 and LOV2) domains at the N-terminus, and a linker region that connects the LOV2 and the kinase domains.1,2 Each LOV domain noncovalently binds a flavin mononucleotide (FMN) as a chromophore. The LOV2 domain plays a key role in the regulation of the kinase activity, whereas the proposed functional role of the LOV1 domain is to regulate the light sensitivity of phototropin.8−12 For understanding the molecular mechanism of the biological function, the reactions of the LOV domains have been recently attracting significant attention.13−18 Upon photoexcitation of the FMN (the ground state protein; D447), the triplet state (L660) is created, and the isoalloxazine ring of the FMN makes a covalent bond to a cysteine residue in the LOV domain (S390).16,17 Subsequent conformational changes of the LOV domain and the linker domain are considered to be important for regulating kinase activity. An NMR study showed that the α-helix in the linker region at the C-terminus of the LOV2 domain (Jα helix), which docks onto the β-strands of the LOV2 domain via hydrophobic interactions in the dark state, dissociates from the LOV2 domain upon light illumination.15 The reaction dynamics of the LOV2 domain and LOV2-linker have been studied by the pulse laser induced transient grating (TG) method.19,20 Following photoexcitation of the FMN, the Jα helix dissociates from the LOV2 domain with a time constant of 300 μs (T390pre), and subsequently, the Jα helix is destabilized and unfolded with a time constant of 1 © 2013 American Chemical Society

ms (T390) for the Arabidopsis phot1 LOV2-linker sample. For the phot2 LOV2-linker sample, a subsequent small volume expansion process with a time constant of 11 ms was observed (T390(II)) after the unfolding reaction of the linker region. The T390(II) state returns back to the ground state with a time constant of ≈5 s.21 FT-IR at low temperatures22−24 and optical rotatory dispersion experiments25 also indicated the conformation (unfolding) change in the Jα helix region. It was suggested that the signal from the LOV core is propagated through the Iβ strand to the peripheral Jα helix.27 Biochemical experiments on phototropin from Arabidopsis thaliana have shown that a mutant (I608E) in which the Jα helix was disrupted in the dark did not provide the light-dependent activation of the kinase domain, and the kinase was constitutively active in both dark and light states.23,28 It is widely considered that the conformational change of the Jα helix is vital for kinase activity. However, recently, the importance of another helix located at the N-terminus (A′α helix) was suggested as the signaling route from the LOV2 domain to the kinase domain.26,29 Similar to the Jα helix, this A′α helix has amphipathic character and binds the LOV2 domain via hydrophobic interactions in the dark state.29 Sequence alignment has shown that the A′α helix is highly conserved in a variety of LOV domains.29,30 A crystallographic study of the Avena phot1LOV2-linker revealed light dependent conformational changes in the N- and Cterminal flanking regions.29 Fourier transform infrared (FTIR) Special Issue: Michael D. Fayer Festschrift Received: June 20, 2013 Revised: August 4, 2013 Published: August 9, 2013 15606

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spectroscopy results suggested a conformational change in the A′α helix.22,31 Aihara et al. have shown, using a hybrid assay and an in vitro phosphorylation assay, that point mutations within the A′α helix (e.g., T207I mutant) of phototropin from Chlamydomonas reinhardtii reduced the kinase repressive activity of the LOV2 domain in the dark state.30 More recently, MD simulations predicted that the presence of the A′α helix was necessary to stabilize the Jα helix, while the removal of Jα helix did not affect the stability of the A′α helix.26 These findings suggested that the A′α helix interacts with Jα helix in the dark. Moreover, a residue-level network analysis of the results from the MD simulations indicated that the interaction between the A′α helix and Jα helix was disrupted upon photoexcitation, which might trigger the dissociation of the Jα helix from the LOV2 domain.32 On the basis of these results, it is now suggested that the S390 state formation triggers the A′α helix unfolding first and this change induces the dissociation of the Jα helix. This is an intriguing proposal for the signaling route of the LOV2-linker. However, the conformational change dynamics of the A′α helix should be examined experimentally at physiological temperature in solution. In this study, we investigated the signaling pathway by monitoring the reaction dynamics of wild-type (WT) and two mutants of the phot1LOV2-linker from Arabidopsis. The reaction was detected by the time-resolved TG method, which is a sensitive and powerful method to detect conformation changes of helices in the time-domain.33,34 We chose a phot1LOV2-linker construct containing both the A′α and Jα helices (449I−661K) as the WT protein and mutants I608E and T469I. Ile608 is a key residue involved in the hydrophobic interaction between the LOV2 domain and the Jα helix. By replacing this residue, NMR and FTIR analyses showed that the Jα helix is dissociated from the LOV2 domain and unfolded in the dark.23,28 The A′α helix is a very short helix and its helical structure is formed by hydrogen bonding among Thr469, Leu470, and Arg472 residues. One of the hydrogen bonds is lost in the T469I mutant, and the helix is considered to be unfolded. This unfolding was confirmed by CD analysis in the present study before examining the reaction dynamics. In addition to these mutants, the helical contents of two other mutantsthe ΔA′α mutant, which lacks the A′α helix (474E− 661K), and the ΔA′α/ΔJα mutant, which lacks both the A′α and Jα helices (474E−586R)were investigated in the dark. By comparing the conformational changes that occur to the mutants and the WT protein, the relationship between the A′α helix and Jα helix was revealed.

After centrifugation, the supernatant was purified by the GSTrap column. GST tags were removed by PreScission Protease digestion. The cleaved polypeptides were purified further by gel chromatography with Sephacryl S-100 HR (GE Healthcare) and eluted with the PBS buffer solution. Circular Dichroism Spectroscopy. The secondary structures of the mutants in the ground state were examined by circular dichroism (CD) spectroscopy (J-720WI, JASCO). The concentrations of the samples were determined by the Bradford protein assay. We also measured the recovery of secondary structural changes induced by light illumination. In this experiment, the lit state was accumulated by illumination with a blue LED (472 nm, 3W) for 3 s, and over 98% of the protein molecules were converted to the lit state under our experimental conditions. The blue light was then turned off, resulting in the characteristic structural recovery of the dark state, which was monitored as a recovery of the CD intensity at 222 nm. During the measurements, samples were stirred with a magnetic stirrer, and the temperature was 23 °C. Transient Grating Method. A laser pulse from a dye laser (Lumonic, HyperDye 300; wavelength, 462 nm) pumped by an excimer laser (Lambda Physic, XeCl operation; 308 nm) was used as a pump beam. 39 The repetition rate of the photoexcitation was 0.01 Hz. This interval between the excitation pulses is sufficiently longer than the lifetime of the product. A diode laser (835 nm) was used as a probe beam. Measurement of the I608E mutant was carried out at a relatively low concentrations (25 μM), since this sample tended to aggregate at high concentrations. In the case of T469I mutant, the measurement was carried out at a concentration of 100 μM. The concentrations of the samples were determined by absorbance at 447 nm using the extinction coefficient of 14300 M−1 cm−1. All measurements were carried out at 23 °C.

EXPERIMENTAL METHODS Preparation of Samples. A DNA fragment encoding the A. thaliana phot1LOV2-linker (residue 449E−661K) and phot1LOV2 (residue 449E−586R) were cloned into the pGEX6P-1 vector. All mutants were prepared by using the plasmid as a template and the PrimeSTAR Mutagenesis Basal Kit (TaKaRa). The I608E and T469I mutants were designed by according to previous work.23,28,30 Primers used for these mutagenesis steps are listed in Table 1. Escherichia coli strain JM109 was transformed by the expression vectors and the cell was grown at 37 °C until the cell density (OD600) reached 0.9. IPTG was added to a final concentration of 0.1 mM, and the cells were incubated for a further 16 h at 20 °C. Harvested bacteria were lysed in phosphate-buffered saline (PBS) containing 140 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, and 1.8 mM KH2PO4 (pH 7.5).

RESULTS CD Measurements. The CD spectra of WT and mutants of the LOV2-linker in the dark state are shown in Figure 1. The negative intensity at 222 nm, which predominantly represents the amount of helical structure, was strongly affected by the mutations. The spectra of the I608E mutant showed a significant loss of helical structure, which was attributed to the unfolding of the Jα helix, as reported before.23,28 On the other hand, the T469I mutant showed a very small change in secondary structure. We did not expected this small change, because, according to the previous MD simulations result, the Jα helix is destabilized by this mutation and the A′α helix unfolds.26 On the contrary, this small change is reasonable, if we consider that the unfolding of the A′α helix does not affect the stability of the Jα helix, because the A′α helix consists of only four residues.

Table 1. List of Primers Used in This Study to Make Several Derivatives of the phot1LOV2 Domain from Arabidopsis thaliana mutant T469I I608E ΔA′α and ΔA′α/ΔJα

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primer 5′-GCTACTATTCTCGAACGTATCGAGAAG-3′ 5′-TTCGAGAATAGTAGCTAGATCAATACC-3′ 5′-GTGAATGAGGATGAAGCGGTTCGAGAACTT-3′ 5′-TTCATCCTCATTCACAGCTGTTTTTTTCAC-3′ 5′-GGAATTCGAGAAGAATTTCGTCATC-3′ 5′-TTCTTCTCGAATTCCGGGGATCCCAG-3′

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Figure 1. CD spectra of the phot1LOV2 derivatives in the dark state. WT, T469I, ΔA′α, ΔA′α/ΔJα, and I608E are shown as solid, chain, broken, circle, and dotted lines, respectively.

Figure 2. Temporal profiles of the CD intensities at 222 nm after terminating the light illumination for: (a) ΔA′α/ΔJα, (b) I608E, and (c) LOV2 linker (dotted lines). The best fitted curves by a single exponential function are shown by the solid lines. These profiles represent the recovery dynamics of the secondary structure.

For further confirmation of this unexpected result, we measured the CD spectrum of a mutant, in which the A′α helix was removed (474E−661K: ΔA′α mutant). The absolute CD intensity at 222 nm was weaker than the value for the T469I construct. Although the structure of the LOV2 domain might be changed by the removal of A′α helix, we excluded this possibility based on the observation that the crystal structure of the LOV2 domain was unchanged in the absence of the Nterminal helical region.26 Instead, we consider that this result indicates a partial unfolding of the Jα helix by the elimination of the A′α helix. These results indicate that the Jα helix indeed interacts with A′α region but the helical structure of A′α is not necessary to stabilize the Jα helix. A crystallographic analysis reported that Lys475, located between the A′α helix and the LOV2 core, and Thr604, present in the Jα helix, formed a hydrogen bond.29 This noncovalent interaction may be important in stabilizing the helical structure of the Jα helix. Therefore, when the A′α helix was removed, the position of Lys475 might be shifted away from Thr604, which destabilizes the Jα helix. The fact that the disruption of the A′α helix did not affect the stability of Jα helix indicated that the presence of the A′α region (regardless of fold) is sufficient to maintain the position of Lys475 and, therefore, to stabilize the Jα helix. When both the A′α and Jα helices were removed (474E−586R: ΔA′α/ΔJα mutant), the absolute CD intensity of this sample was much smaller than the other samples. This is reasonable, because the very long structure (99 residues) containing the αhelix was removed.35,36 Next, the secondary structure under the light condition was investigated by monitoring the light induced change of the CD intensity at 222 nm. Figure 2 shows the recovery process of the CD intensities at 222 nm of several mutants. The signals were well-reproduced by a single-exponential function, and the lifetimes of the lit state were almost identical to each other and determined to be 60 ± 5 s. This lifetime was very similar to the recovery process monitored by the absorption detection of the chromophore (55 ± 5 s at 23 °C). This observation indicates that the secondary structure was recovered in synchrony with the disruption of the covalent bond between the chromophore and the protein. The amount of the secondary structural change upon light illumination was evaluated from the amplitude at t = 0 of the exponential curve. The amplitudes were 1.8 mdeg for ΔA′α/ΔJα, 2.0 mdeg for I608E, and 8.7 mdeg for WT. The value of the WT protein was much larger than those of the other two samples. This large change was attributed to the

unfolding of both A′α and Jα helices upon light illumination. The ΔA′α/ΔJα also showed a small change in its secondary structure, despite lacking both helices. We attributed this change to a distortion of the β-strands of the LOV2 domain, because the crystallographic and NMR studies have reported that the helical structure was not unfolded in the LOV2 domain, and instead, small conformational changes in β-strands were observed upon light illumination.15,29 By comparing the amplitude of secondary structural change between ΔA′α/ΔJα and I608E, we found that the I608E showed a slightly larger change. We consider that this small difference reflects the unfolding of the A′α helix upon light illumination. TG Measurement. The kinetics of the conformation change of the A′α helix was investigated by the time-resolved TG method. In the TG method, the photochemical reaction is induced by an interference pattern of two excitation beams, which leads to refractive index modulations.37−39 A probe beam is diffracted by this refractive index grating, and the signal intensity is proportional to the square of the refractive index change (δn). A typical TG signal of T469I (A′α helix unfolded mutant) at a concentration of 100 μM and at a square of the grating wavenumber q2 = 5.3 × 1010 m−2 is shown in Figure 3. The signal rose quickly within the pulse width of the excitation laser, and the signal decayed in a submicrosecond time range followed by two rise-decay components before finally decaying

Figure 3. An example of the observed TG signal (dotted line) of T469I at a concentration of 100 μM at q2 = 5.3 × 1010 m−2. The best fitted curve based on the two-state model is shown by the solid line. 15608

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function, of which the rate constants are given by the diffusion coefficients of the reactant (DR) and the product (DP), times, and q2:

to the baseline. This overall qualitative temporal feature of the signal is similar to that of the WT protein reported previously.20 The signal can be decomposed into several components, and the assignment of the kinetics are described as follows. The initial decay-rise signal can be reproduced well by a single exponential function with a time constant of 1.9 μs, which corresponds to the kinetics of covalent bond formation between the chromophore and a cysteine residue (the adduct (S390) formation process). The next decay rate was dependent on the q2 value. By comparing this with the thermal grating signal from a calorimetric sample (bromocresol purple in aqueous solution), it was concluded that this signal is the thermal grating signal, which is created by the thermal energy due to the nonradiative transition from the excited state, and it decayed with a rate constant of Dthq2 (Dth: thermal diffusivity of the solution, q: grating wavenumber). The rise and decay profile appearing after this thermal diffusion signal is the species grating signal (δnspe(t)), which reflects the chemical reaction kinetics as well as the molecular diffusion process. Therefore, the total TG signal (ITG(t)) is expressed as:

ITG(t ) = α{δnP0 exp(−DPq2t ) − δnR0 exp(−DR q2t )}2

δn0R(>0)

where and are, respectively, the initial refractive index changes due to the changes in the reactant and the product concentrations. Therefore, the peak intensity of the diffusion signal depends on the difference between the DP and DR values. If the difference is larger, the peak intensity is larger. The similar signal intensity of T469I to that of the WT indicates that the difference is similar. Since the sign of the refractive index of the thermal grating is negative (δnth < 0) at this temperature, we determined the signs of the refractive index change of the rise and decay components to be negative and positive, respectively. From these signs, the rise and decay components of the diffusion signal are attributed to the protein diffusion processes of the reactant (ground state protein) and the photoproduct, respectively. The faster rate of the rising component than that of the decay indicates that the product diffuses slower than the reactant (DR > DP). Using a curve fitting routine, we determined DR and DP of the WT protein under this condition to be 9.2 × 10−11 m2/s and 6.2 × 10−11 m2/s, respectively.40 DR and DP of T469I were 8.9 × 10−11 m2/s and 6.2 × 10−11 m2/s, respectively. The diffusion signals measured at various q2 are depicted in Figure 5. The signals were normalized with the number of

ITG(t ) = α{δna exp( −ka t) + δnth exp( −Dth q2t ) + δnspe(t )}2

(2)

δnP0(>0)

(1)

where α is a constant, ka is the rate constant of the adduct formation, and the pre-exponential factors are the refractive index changes by these processes. Hereafter, the time profile of only δnspe(t) is analyzed, because the other part of the signals of the mutants are almost identical to those of the WT species, which have been described previously.19,20 To determine the kinetics of the α-helices unfolding process, we first determined the diffusion coefficients of the reactant (DR) and the final product (DP). For that purpose, we measured the TG signal at a relatively small q2 (5.3 × 1010 m−2). At this q2, the diffusion peak appeared in a time range of 0.1−1 s. This time range is sufficiently slower than the reaction rate (vide infra), so that a component representing the reaction kinetics did not contribute to the signal. For comparison, we also measured the TG signals of the WT and I608E (Jα helix unfolded mutant) mutant under identical experimental conditions, and the results are depicted in Figure 4. The diffusion signal intensity of T469I was similar to that of the WT. When we ignore the reaction kinetics, the temporal profile of the species grating signal (ITG(t)) is expressed by the diffusion of the reactant and product, hence, by a biexponential

Figure 5. q2 dependence of the TG signal (dotted line) of T469I at a concentration of 100 μM. The q2 values are: (a) 5.3 × 1012; (b) 3.4 × 1012; (c) 1.9 × 1012; (d) 4.8 × 1011; and (e) 1.5 × 1011 m−2. The signals representing the molecular diffusions processes are shown, and these signals are normalized with the number of excited molecules. The best fitted curves by the two-state model are shown by the solid lines. The signals are closely matched by the simulated fitted curves.

reacted molecules, which was determined by the thermal grating signal intensity. The intensity of the diffusion peak of the T469I mutant strongly depended on the grating wavenumber (Figure 5). This time dependence of the signal intensity can be explained in terms of the time-dependent D change. On a short time scale, the diffusion signal intensity was weak, because the change in D for the product is small at this time, and the two terms in eq 2 canceled each other. With increasing time, DP gradually decreased and the difference between DP and DR increased, so that the diffusion peak intensity increased. This time-dependent D change was analyzed by using the following model:

Figure 4. TG signals of (a) WT, (b) T469I, and (c) I608E at a concentration of 25 μM and q2 = 4.0 × 1010 m−2. 15609

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k

R→I→P

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The diffusion signal of I608E measured at q2 = 4.0 × 10−10 m was much weaker than that of the WT and also that of T469I (Figure 4). This weak signal intensity is explained in terms of a small D change by the reaction. Indeed, we determined DP and DR by biexponential fitting of the signal to be DR = 6.6 × 10−11 m2/s and DP = 6.2 × 10−11 m2/s. The D change of I608E associated with the unfolding of A′α helix was much smaller than that of WT (DR = 9.2 × 10−11 m2/s, DP = 6.2 × 10−11 m2/s). This is reasonable because only the short A′α helix unfolds in the I608E mutant, as shown in the CD measurement (Figure 2). The diffusion signal intensity normalized with the number of excited molecules depended on q2; the signal intensity increased as the observation time range increased (Figure 6).

(Scheme 1)

−2

where R, I, P, and k represent respectively, the reactant (D447), an intermediate species (S390), a final product (T390), and the rate constant of the change. Solving a diffusion equation, the temporal profile of the diffusion signal created by the above reaction is obtained as: ⎞ ⎛ k ITG(t ) = [⎜δnI − δnP⎟ 2 (D I − D P )q + k ⎠ ⎝ exp{− (DIq2 + k)t } k δnPexp( −DPq2t ) + (D I − D P )q 2 + k − δnR exp(−DR q2t )]2

(3)

where δnI and DI are the refractive index change due to the creation of the intermediate species and the diffusion coefficient of the intermediate species, respectively. Using DR and DP determined above, and the observed TG signals were reproduced reasonably well over a wide observable time range (Figure 5). The diffusion coefficient of the intermediate and the time constant of the reaction were determined to be 8.1 × 10−11 m2/s and 1.0 ms, respectively. The origin of the change in D by a chemical reaction has been classified into two categories: oligomer formation and conformational change (i.e., diffusion-sensitive conformation change).33,35,41,42 We determined the origin by the concentration dependence of the rate. If the D change is caused by an oligomer formation process, the kinetics of the TG signal should be sensitive to the concentration.42 However, the temporal profile of the diffusion signal (except the absolute intensity) was independent of the protein concentration (data not shown). Therefore, we concluded that the D change is due to an intramolecular conformational change. This origin is the same as that for the WT protein reported previously. When the Jα helix unfolds, hydrophilic residues that stabilize the helical structure in the dark state are exposed to the solvent and may enhance the interaction with the solvent molecules. This change leads to the observed decrease in D. The CD spectrum (Figure 1) showed that the T469I mutant kept the helical structure of Jα in the dark state and the helix unfolds upon light illumination. Hence, the D change should be caused by an enhancement of the intermolecular interactions between the protein and the solvent associated with the unfolding process. The difference between DP and DR was similar to that of the WT. This indicates that the D change of the WT is mainly attributed to the unfolding of the Jα helix and the contribution of the unfolding of the A′α helix was minor. This is consistent with the results of the CD experiments. The slightly smaller DR of T469I than that of the WT is reasonable, because the A′α helix of the reactant of T469I was unfolded and the unfolding of the α-helix causes the decrease in D owing to an increase in friction.33,34 Almost the same DP between WT and T469I suggests that the product is the same; that is, both of the Jα and A′α helices are unfolded in the product (vide infra). Interestingly, the reaction time constant of the unfolding process of the Jα helix in the T469I mutant was identical to that of the WT (1 ms), which suggests that the reaction kinetics of the Jα helix were conserved, even when the A′α helix was unfolded in the dark.

Figure 6. TG signals (broken lines) of the I608E mutant at q2 values of: (a) 3.9 × 1012; (b) 6.5 × 1011; (c) 1.6 × 1011; and (d) 3.7 × 1010 m−2. The signals representing the molecular diffusion processes are shown, and these signals are normalized with the number of excited molecules. The best-fitted curves to the observed TG signals by the two-state model (eqs 1 and 3) are shown by the solid lines. The signals are closely matched by the simulated fitted curves.

This indicated that DP was time-dependent in the observation time range (100 μs to 20 ms). Using DP and DR determined above, the diffusion coefficients of the intermediate and the time constant of the reaction were determined to be 6.5 × 10−11 m2/s and 12 ms, respectively.

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DISCUSSION Previous studies have suggested that the interaction between the A′α and Jα helices plays a key role in regulating signal transduction. The interaction changes upon light illumination with the A′α and Jα helices dissociating from the LOV2 domain to promote signaling.26,32 In this study, we examined the interaction and the reaction kinetics of these helices by using the CD and TG methods. The CD measurements proved that the existence of the A′α helix was important to stabilize the Jα helix (the Jα helix was partially unfolded in the ΔA′α mutant). However, the helical structure of the A′α is not necessary for the stabilization (the Jα helix was not unfolded in the T469I mutant). Hence, we postulate that the contact (or the hydrophobic interaction) between the A′α and the Jα helices is not essential to maintain their helical structures in the dark state, but the hydrogen bond between Lys475 and Thr604 is important. According to the crystallographic structure, the A′α and the Jα helices are located in close proximity to each other, and it was suggested that the hydrophobic interactions between these two helices exist.29 On the other hand, the solution NMR structure showed that the Jα helix is bent between residues Lys602 and Ala605.29 This structure suggests that the 15610

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phot1LOV2 without the Jα helix showed a monomer−dimer equilibrium in the dark state, and the reaction scheme showed the dimerization and dissociation reaction.19 The small conformational change of the A′α helix was difficult to detect. However, in the case of the Arabidopsis phot2LOV2 construct, which includes both the A′α helix and the Jα helix, a small volume change was observed previously with a time constant of 11 ms, and this was in addition to the unfolding of the Jα helix.21 This signal was also observed for the phot2LOV2 construct, which contains only the A′α helix. At that time, we could not assign this dynamic process. However, the similar time constant (12 ms for the A′α helix unfolding vs 11 ms volume change) suggests that the small volume change might represent the unfolding of A′α helix. If the unfolding of the A′α and Jα helices of phot1LOV2 occurs independently, the reaction scheme for LOV2-linker may be described by the following scheme:

interaction between these helices cannot be strong. Thus, our observation is consistent with the NMR structure. Previously, it was reported that the Jα helix unfolds when a single mutation is introduced (I608E). This mutation may weaken the hydrophobic interaction between the LOV2 domain and the Jα helix.28 Therefore, the Jα helix is stabilized by the interaction with the β-strands of the LOV core, and the presence of the A′α helix is not crucial. From the TG measurement of the I608E mutant, we found that the A′α helix unfolded with a time constant of 12 ms, which is slower than the unfolding of the Jα helix of the WT protein (1 ms). Hence, we consider that the previously proposed signaling route;26 that is, the unfolding of the Jα helix is triggered by the unfolding of the A′α helix, cannot be possible. Moreover, it is interesting to note that the T469I mutant, which lacks the helical structure of the A′α helix, showed the same reaction dynamics as that of the WT. These findings suggest that conformational changes of the A′α and Jα helices occurred independently. The schematic illustration of the conformational change process is shown in Figure 7.

where T390(I) and T′390(I) represent the unfolded states of the Jα helix and the A′α helix, respectively. However, in this case, since the rate of the A′α helix unfolding is much (10 times) slower than that of the Jα helix, most of the reaction proceeds by the upper branch of the above scheme, that is, the A′α helix unfolding process occurs after the Jα helix unfolding. Therefore, the reaction scheme may be simplified as hν

Figure 7. Schematic showing the photoreaction of the phot1LOV2linker containing the A′α helix.

1.9μs

ns

300μs

1ms

pre D447 → D*447 → L660 ⎯⎯⎯⎯→ S390 ⎯⎯⎯⎯⎯→ T390 ⎯⎯→ T390(I) 12ms

∼ 70s

⎯⎯⎯⎯→ T390(II) ⎯⎯⎯⎯→ D447

where T390(I) is the unfolded state of the Jα helix and T390(I) → T390(II) corresponds to the conformational change of the A′α helix. In this study, we suggest that the unfolding reactions of the A′α and Jα helices occur independently. Additionally, it has been suggested that a mutation on either the A′α helix or the Jα helix inhibits the light-dependent regulation of the kinase activity (kinase domain is constitutively active in both the dark and light states).28,30 Taking these findings into account, we consider that the kinase domain would be activated when one of these helices are disrupted and the signal transfer from the LOV2 domain to these helices might be achieved. Currently, it is postulated that a structural change to the Jα helix is vital for the activation of the kinase; however, this concept has been challenged by Matsuoka and Tokutomi.10 They showed that kinase activity was regulated in a light-dependent manner when the isolated kinase domain was mixed with the LOV2 domain that lacks the Jα helix.10 This observation indicates that the Jα helix is not involved in the regulation of kinase activity. Since the LOV2 construct used in that report contains the A′α helix, we hypothesize that the A′α helix as well as the Jα helix regulates kinase activity. It was proposed that the kinase activity is suppressed by the docking of the LOV2 domain, and the release of the LOV2 domain by light illumination changes interdomain interactions between the LOV2 domain and the kinase domain, thereby leading to the activation of the

If helices are composed of the same residues, a longer helix may be more stable than a smaller one. Nevertheless, the A′α helix unfolds slower than the Jα. This different unfolding rates between the Jα and A′α helix may reflect the different stabilities of the helices as the polypeptide chains. Indeed, a disorder probability calculation showed that the sequences of the Jα and A′α helices were classified as naturally unfolded polypeptides that adopt several conformations, dependent on the interactions with other polypeptide chains.44 Thus, when the Jα helix dissociates from the β sheet of the LOV2 domain, the Jα helix readily unfolds. Similarly, the A′α helix is also unstable in nature, but the stability is slightly higher than that of the Jα helix.44 This higher stability could cause the slower rate of the reaction. Moreover, the Jα helix is located near the Hβ-Iβ loop region, which was shown to fluctuate highly by the MD simulations.45 This larger fluctuation may cause the faster unfolding of the Jα helix. In our previous study on phot1LOV2 containing both the A′α helix and the Jα helix, the signal of the unfolding of the A′α helix could not be detected.20 This is not surprising, because the A′α helix is very short (only four residues) and the signal arising from this helix is weak and is masked by the strong diffusion signal representing the unfolding of the Jα helix. In fact, the molecular diffusion signal of I608E associated with the unfolding of A′α helix was negligibly small when compared with that of the WT protein (Figure 4). Furthermore, 15611

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kinase.43,44 According to SAXS data, the structure of the LOV2linker-kinase elongates following light illumination.44 The restored molecular models indicated that the centers of the LOV2 and kinase domains were separated in the light state. When we fit the LOV2-linker domain in the restored model, we found that the space between the LOV2 and kinase domains corresponds to the Jα and A′α helices. Thus, it may be reasonable to consider that the conformational change of the A′α helix also regulates the separation of the LOV2 domain from the kinase domain. Interestingly, a previous study by FTIR reported that the conformational change of the Jα helix and the interaction between the LOV-core and the Jα helix were variant and dependent on phototropins.24 On the other hand, the amino acid sequence of the A′α helix is better conserved than that of the Jα helix. From these points of view, we consider that the A′α helix provides robust regulation of the kinase activity, and the Jα helix yields a variety of mechanisms of signal transduction.

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(10) Matsuoka, D.; Tokutomi, S. Blue Light-Regulated Molecular Switch of Ser/Thr Kinase in Phototropin. Proc. Natl. Acad. Sci. U.S.A. 2005, 102 (37), 13337−13342. (11) Salomon, M.; Lempert, U.; Rudiger, W. Dimerization of the Plant Photoreceptor Phototropin Is Probably Mediated by the LOV1 Domain. FEBS Lett. 2004, 572 (1−3), 8−10. (12) 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. (13) Moglich, A.; Yang, X. J.; Ayers, R. A.; Moffat, K. Structure and Function of Plant Photoreceptors. Ann. Rev. Plant. Biol. 2010, No. 61, 21−47. (14) Herrou, J.; Crosson, S. Function, Structure and Mechanism of Bacterial Photosensory LOV Proteins. Nat. Rev. Microbiol. 2011, 9 (10), 713−723. (15) Harper, S. M.; Neil, L. C.; Gardner, K. H. Structural Basis of a Phototropin Light Switch. Science 2003, 301 (5639), 1541−1544. (16) Schuttrigkeit, T. A.; Kompa, C. K.; Salomon, M.; Rudiger, W.; Michel-Beyerle, M. E. Primary Photophysics of the FMN Binding LOV2 Domain of the Plant Blue Light Receptor Phototropin of Avena Sativa. Chem. Phys. 2003, 294 (3), 501−508. (17) Kennis, J. T. M.; Crosson, S.; Gauden, M.; van Stokkum, I. H. M.; Moffat, K.; van Grondelle, R. Primary Reactions of the LOV2 Domain of Phototropin, A Plant Blue-Light Photoreceptor. Biochemistry 2003, 42 (12), 3385−3392. (18) Schleicher, E.; Kowalczyk, R. M.; Kay, C. W. M.; Hegemann, P.; Bacher, A.; Fischer, M.; Bittl, R.; Richter, G.; Weber, S. On the Reaction Mechanism of Adduct Formation in LOV Domains of the Plant Blue-Light Receptor Phototropin. J. Am. Chem. Soc. 2004, 126 (35), 11067−11076. (19) 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. (20) 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. (21) Eitoku, T.; Nakasone, Y.; Zikihara, K.; Matsuoka, D.; Tokutomi, S.; Terazima, M. Photochemical intermediates of Arabidopsis Phototropin 2 LOV Domains Associated with Conformational Changes. J. Mol. Biol. 2007, 371 (5), 1290−1303. (22) Alexandre, M. T. A.; van Grondelle, R.; Hellingwerf, K. J.; Kennis, J. T. M. Conformational Heterogeneity and Propagation of Structural Changes in the LOV2/J alpha Domain from Avena sativa Phototropin 1 as Recorded by Temperature-Dependent FTIR Spectroscopy. Biophys. J. 2009, 97 (1), 238−247. (23) Yamamoto, A.; Iwata, T.; Sato, Y.; Matsuoka, D.; Tokutomi, S.; Kandori, H. Light Signal Transduction Pathway from Flavin Chromophore to the J alpha Helix of Arabidopsis Phototropin1. Biophys. J. 2009, 96 (7), 2771−2778. (24) Koyama, T.; Iwata, T.; Yamamoto, A.; Sato, Y.; Matsuoka, D.; Tokutomi, S.; Kandori, H. Different Role of the J alpha Helix in the Light-Induced Activation of the LOV2 Domains in Various Phototropins. Biochemistry 2009, 48 (32), 7621−7628. (25) Chen, E. F.; Swartz, T. E.; Bogomolni, R. A.; Kliger, D. S. A LOV Story: The Signaling State of the Phot1 LOV2 Photocycle Involves Chromophore-Triggered Protein Structure Relaxation, As Probed by Far-UV Time-Resolved Optical Rotatory Dispersion Spectroscopy. Biochemistry 2007, 46 (15), 4619−4624. (26) Zayner, J. P.; Antoniou, C.; Sosnick, T. R. The Amino-Terminal Helix Modulates Light-Activated Conformational Changes in AsLOV2. J. Mol. Biol. 2012, 419 (1−2), 61−74. (27) Nash, A. I.; Ko, W. H.; Harper, S. M.; Gardner, K. H. A Conserved Glutamine Plays a Central Role in LOV Domain Signal Transmission and Its Duration. Biochemistry 2008, 47 (52), 13842− 13849.

AUTHOR INFORMATION

Corresponding Author

*Tel./Fax: +81-75-753-4026; e-mail: mterazima@kuchem. kyoto-u.ac.jp. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by Grant-in-aid for Scientific Research on Innovative Areas (research in a proposed research area) (20107003) from the Ministry of Education, Culture, Sports, Science and Technology in Japan (to M.T.).

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REFERENCES

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