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Feb 12, 2018 - Mechanistic Basis of the Fast Dark Recovery of the Short LOV Protein ... Photoreaction Dynamics of LOV1 and LOV2 of Phototropin from ...
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Article Cite This: Biochemistry XXXX, XXX, XXX−XXX

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Sequential DNA Binding and Dimerization Processes of the Photosensory Protein EL222 Akira Takakado, Yusuke Nakasone, and Masahide Terazima* Department of Chemistry, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan S Supporting Information *

ABSTRACT: EL222 is a blue light sensor protein, which consists of a light− oxygen−voltage domain as a light sensor and a LuxR-type helix−turn−helix DNAbinding domain. The reaction dynamics of the protein−DNA binding were observed for the first time using the time-resolved transient grating method. The reaction scheme was determined, showing that photoexcited EL222 first binds DNA and the ground state EL222 monomer is subsequently associated with the complex. Rate constants on the millisecond scale were determined for these processes. In addition, binding rates for EL222 with three DNA sequences, with different binding affinities, were measured. Although EL222 binds nonspecific DNA sequences with affinities at least 5-fold lower than the target sequence affinity, the binding rates were almost the same as that for the target DNA. This observation indicates that the specific and nonspecific binding affinities are mainly controlled by differences in the dissociation of DNA binding.

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rotein−DNA interaction is a fundamental process for DNA expression and has been studied using various methods such as footprinting assays, electrophoretic mobility shift assays (EMSAs), and surface plasmon resonance (SPR).1,2 However, kinetic information for the process has been limited, because of experimental difficulties, for example, problems with initiating the reaction and detecting the dynamics.3 With regard to these points, photosensory proteins that can be triggered by light are useful for studying the dynamics. In this study, we investigated the reaction dynamics of protein−DNA interaction for a photosensory DNA-binding protein, EL222. EL222 is a blue light sensory protein from the bacterium Erythrobacter litolaris HTCC2594.4 It consists of a light− oxygen−voltage (LOV) domain, as a light-sensing domain in the N-terminal half, and a LuxR-type helix−turn−helix (HTH) DNA-binding domain in the C-terminal half, as an effector domain.4,5 In the dark, EL222 shows negligible interaction with DNA, while upon light activation, DNA binding is significantly enhanced.4,6 The DNA sequence of the EL222-binding motif was shown to be 5′-RGNCYWWRGNCY-3′ (Y = C or T, W = A or T, R = A or G, and N = any nucleotide).6 A biochemical analysis reported that EL222 is a light-dependent transcriptional factor.6 This EL222−DNA interaction has been attracting significant attention, not only because protein− DNA interaction is important for understanding DNA expression but also because EL222 has been used for robust regulation of gene expression as a useful tool in optogenetics.7−9 The LOV domain noncovalently binds a flavin mononucleotide (FMN) as a chromophore when the domain is in the dark. Upon light illumination, it forms a covalent adduct with a nearby cysteine residue, which triggers conformational changes in the protein, transmitting a light signal.10−12 It has been reported that EL222 exists as a monomer in the dark.4,13 © XXXX American Chemical Society

However, because typical LuxR-type HTH proteins bind to DNA as dimers, it is reasonable to think that EL222 may interact with DNA as a dimeric form.4,14 Indeed, size exclusion chromatography (SEC) and EMSA experiments have shown that two EL222 proteins associate with DNA in the illuminated state.4,14 Light-induced dimerization therefore appears to be necessary for the activation. However, it is not clear how the monomer in the dark state forms the dimeric EL222−DNA complex by light illumination. Clarifying the intermolecular interaction change was considered a useful starting point for elucidating the reaction mechanism. Our previous study using the time-resolved transient grating (TG) method showed that the LOV domain of EL222 (EL222-LOV domain) is in equilibrium between the monomer and dimer in the dark, and it exhibits light-induced dimerization very efficiently. On the other hand, the dimerization yield of full-length EL222 is very low, although the photoexcitation of the monomer leads to dimerization with a rate constant of 8.6 × 103 M−1 s−1.13 Our findings and the crystal structure in the dark suggest that the LOV domain and 4α-helix in the HTH domain are the dimerization sites, and these domains are blocked by the interacting HTH domain in the dark state.4 Light illumination should therefore induce the dissociation of the LOV and HTH domains, which in turn should be relevant for the dimerization.4 We investigated the DNA binding dynamics of EL222 using the time-resolved TG method, which can detect intermolecular interaction processes as changes in the diffusion coefficient. We successfully determined the reaction scheme of DNA binding, Received: November 30, 2017 Revised: February 8, 2018 Published: February 12, 2018 A

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry

Electrophoretic Mobility Shift Assay (EMSA) Measurement. The EMSA experiment was performed under white light illumination from a Xe lamp (MAX 300, Asahi Spectra) or in the dark. All measurements were performed at room temperature (23 °C). To establish a light-activated state, samples were incubated under the Xe lamp for 10 min prior to measurement and continuously illuminated during running. For the dark state measurement, the sample was prepared under dim red light and the measurement was performed in the dark. Samples were separated on a 5 to 20% acrylamide gel in TAE buffer for 1 h (ATTO AE-7300). After being run, gels were stained with EtBr for 2 h and monitored by fluorescence imaging with an excitation wavelength of 308 nm (BioDoc-it UVP imaging system). The concentration of DNA was 0.5 μM, and the concentrations of EL222 were 0, 0.1, 0.5, 1, 2, 5, 10, 20, 40, 60, 80, and 100 μM. Each lane was loaded with 8 μL of sample. Absorption Measurement. The dark recovery rate was measured by the absorption change at 450 nm, which was recorded using an ultraviolet−visible spectrometer (model 8453, Agilent) at 23 °C. The concentration of the proteins and DNA (sequence 1) was 100 μM. The samples were loaded in a quartz cell (optical path length of 2 mm). The activated state was established by blue light illumination from a light-emitting diode (LED) light source (CHR-3S, B3VP-8, Nissin Electronic Co. Ltd.) for 1 min.

as well as the reaction rates for the DNA binding and dimerization processes. We also studied the sequence dependence of the reaction kinetics to understand the molecular mechanisms of DNA sequence recognition. Interestingly, we found that the rate of association with DNA was independent of the DNA sequence, suggesting that the dissociation rate is a dominant factor in determining the binding constant. On the basis of these observations, we proposed a DNA recognition mechanism of EL222.

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MATERIALS AND METHODS Full-length EL222 (residues 1−222) and the LOV fragment (residues 1−144) of EL222 from E. litoralis HTCC2594 were prepared by bacterial overexpression as shown in our previous report.13 The proteins were made in a buffer solution containing 50 mM Tris and 100 mM NaCl (pH 8.0), and all subsequent measurements were performed under these buffer conditions. To investigate the DNA binding dynamics, three DNA fragments were prepared. Sequence 1 was 5′-GGTAGGATCCATCGGGCAGTGCGGTCAGCGGCATGCCGGCAGCAG-3′. This sequence was termed oligomer1 in ref 4. Because the binding affinity was assessed previously using EMSA and nuclear magnetic resonance (NMR) measurements,4,14 we mainly used this sequence for investigations of the reaction dynamics. For comparison, sequence 2 was 5′-TTGCGAGAAGAAAATATGGACCTTGGCCCATGATGGACACAATAC-3′, which was termed peak A in ref 6. Sequence 3 (5′-AAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAA-3′) and poly-dA were used to examine the kinetics of binding DNA with different affinities. All fragments consisted of 45 bp (28 kDa for double-stranded DNA). Single-stranded oligonucleotides for the DNA fragments and their complementary pairs were purchased from Macrogen Japan and annealed by being heated to 95 °C in a heat block for 5 min and then slowly cooled to room temperature. Transient Grating (TG) Measurement. In the TG experiment, two laser beams were crossed at a spot in a sample solution within a coherence time of the light to induce an interference pattern of the light intensity. The intensity is spatially modulated, and molecules in this bright region are photoexcited. If this photoexcitation creates the refractive index grating, a probe beam was diffracted by the grating, and it produces the TG signal. This technique is highly sensitive, because solely photoinduced changes can be detected without contributions from unreacted species. Furthermore, by monitoring the change in D, we detected the dissociation/ association processes sensitively as previously demonstrated for several photosensor proteins.3,13,15,16 The experimental setup of the TG measurement was the same as that described in a previous report.13 A XeCl excimer laser (308 nm, Compex102, Lambda Physik, Fort Lauderdale, FL) pumped dye laser (465 nm, HyperDye 300, Lumonics, Denver, CO) was used as the excitation laser pulse, and a diode laser (835 nm, Crysta Laser, Reno, NV) was used as a probe beam. The TG signal was detected by a photomultiplier tube (R1477, Hamamatsu). In general, 20 signals were averaged for one measurement. Except during light intensity dependence experiments, the signals were measured at a weak laser power not to saturate the signal intensity (Figure S1a). The square of the grating wavenumber (q2) for each experimental setup was calculated from the decay time of the thermal diffusion signal of bromocresol purple in water. The temperature was 23 °C for all measurements.

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RESULTS TG Signals of EL222 with DNA. The reaction of EL222 with DNA (sequence 1) was investigated using the TG method. The binding affinity between EL222 and the DNA (sequence 1) was previously studied using EMSA and NMR measurements.4,14 A typical TG signal after photoexcitation of EL222 at a concentration of 100 μM with the DNA (100 μM) at a q2 of 3.4 × 1010 m−2 is shown in Figure 1.

Figure 1. TG signals of EL222 with DNA (red line) and without DNA (blue line) at a q2 of 3.4 × 1010 m−2. The concentration of EL222 was 100 μM for each sample, and the concentration of the DNA (sequence 1) fragment was 100 μM. The molecular diffusion signal of EL222 without DNA is enlarged in the inset.

For comparison, the TG signal of EL222 at 100 μM without DNA is also shown. These signals are almost identical on the fast time scale (100 ns to 1 ms); i.e., the signals rose quickly within an experimental response time (∼20 ns) and then showed decay−rise and decay components. These phases were assigned to cysteinyl adduct formation of the chromophore in the LOV domain and thermal diffusion.13 The temporal profile of the TG signal in this time range is expressed as B

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry ITG = α[δnad exp( − kadt ) + δnth exp( − Dthq2t ) + δnspe(t )]2

with the DNA. This is consistent with previous studies that demonstrated that EL222 binds to DNA upon light illumination.4,6 Previously, we showed that EL222 exists as a monomer in the dark state and that it dimerizes upon light illumination, although the dimerization yield is low.13 To examine whether the EL222 dimer is involved in the DNA interaction, we investigated the EL222 concentration dependence of the TG signal. Figure 2a shows the TG signals of EL222 at various

(1)

where α is a constant and kad, Dth, and q are the reaction rate constant of adduct formation, the thermal diffusivity of the solution, and the grating wavenumber, respectively. Furthermore, δnad, δnth, and δnspe(t) are the changes in the refractive index caused by adduct formation, thermal grating, and species grating, respectively. Grating wavenumber q is given by 2π/Λ, where Λ is a fringe length of the grating. The sign of the thermal grating is negative at 23 °C (δnth < 0). The rate of adduct formation was determined to be 0.7 μs for both samples. These identical profiles indicate that the DNA does not affect the initial reaction phase of EL222. This observation is consistent with a previous report showing that EL222 does not interact with DNA in the dark state.4 A significant difference between the two samples was observed in the slower time region. Although both signals showed rise and decay components and they were assigned to the molecular diffusion signals, the intensities of the diffusion signals were very different. If reaction dynamics in this time range are ignored, the time profile of the diffusion signal should be expressed by the diffusion of the reactant and the product: δnspe(t ) = −δnR (t ) + δnP(t ) = −δnR exp( −DR q2t ) + δnP exp( −DPq2t )

(2)

where δnR, δnP, DR, and DP are the refractive index changes due to the reactant, the refractive index changes due to the photoproduct, the diffusion coefficient of the reactant, and the diffusion coefficient of the photoproduct, respectively. The prominent rise−decay profile of the signal is a clear indication that the diffusion coefficient is changed by the reaction. From the signs of the refractive index changes, the rise and decay components are assigned to the diffusion of the reactant and the product, respectively. It is apparent that the product diffuses more slowly than the reactant for both samples (DR > DP). If the reaction dynamics are involved in the signal, δnR(t) and δnP(t) can be expressed not only by the diffusion but also by the reaction kinetics and the contribution of intermediate species should be included. The analysis is described in the next section. Previously, the D change of EL222 without DNA was attributed to the photoinduced dimerization reaction and the conformational change associated with the dimerization step.13 The following diffusion coefficients were determined: DR = 8.8 × 10−11 m2 s−1, and DP = 6.6 × 10−11 m2 s−1. Adding DNA to the solution significantly increased the diffusion signal intensity, and the peak position shifted to a slower time. Before quantitatively analyzing the signal, we first qualitatively considered the origin of these changes induced by the presence of the DNA. The observation that the peak time shifted to slower reaction times with the addition of DNA indicates a decrease in the diffusion coefficient. Because the reactant is always EL222, DR should not be changed by the addition of DNA. It is therefore certain that DP decreases as a result of the addition of DNA. Furthermore, the peak intensity of the diffusion signal depends on the difference between DP and DR. The enhancement of the diffusion peak is therefore also rationalized by the decrease in DP, and the increase in the difference between DP and DR. In the next section, we analyze the signals quantitatively, and this explanation is confirmed. The smaller DP indicates interaction of the photoexcited EL222

Figure 2. (a) Protein concentration dependence of the TG signals at a q2 of 4.0 × 1010 m−2. The concentrations of EL222 were 50, 100, 200, 300, 400, and 475 μM, with a constant DNA concentration (sequence 1, 300 μM). The best fit curves using eq S2 are shown as dashed lines. (b) Plot of the reaction rate of the protein dimerization step vs EL222 concentration. The data are fitted with a linear function.

concentrations (50−475 μM) with a DNA concentration of 300 μM. The signals are normalized by the intensity before the diffusion component (∼0.1 ms), which represents the amount of the adduct species. Interestingly, the intensity of the diffusion peak gradually increased with an increase in concentration. This result implies that more than one monomer is involved in the reaction. Later, we show that the EL222 dimer interacts with DNA. Furthermore, we investigated the number of excited EL222 molecules in the dimeric EL222−DNA complex by light intensity dependence. If the dimer in the complex consists of the excited and ground state monomers, this reaction is a onephoton process. On the other hand, if the dimer consists of two excited monomers, the reaction is a two-photon process and the reaction rate should depend on the light intensity.13,15,16 The signals at various light intensities are shown in Figure S1. Even in an unsaturated excitation light intensity range, it was found that the diffusion signal does not depend on the light intensity, when they are normalized by the amount of excited molecules. If the reaction rate changes among these signals, the intensity of the TG signals should be changed. Hence, this result indicates that the reaction rate does not depend on the light intensity. We therefore concluded that dimerization occurs between monomers in the light and dark states. The dimer− DNA complex is therefore composed of one excited and one ground state EL222 molecule [(EL222*−EL222)DNA]. We C

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry previously reported that photoexcited EL222 (EL222*) produces a dimer by interacting with a ground state EL222 monomer, based on similar analysis of the light intensity variation method.13 Our result here indicates that the dimer composition without DNA is the same as that with DNA. Because the excitation light intensity we used was low enough not to saturate the signal (Figure S1a), we consider that the contribution of the dimerization of the two excited EL222 states is small. However, it is difficult to estimate the extents of photoreaction. To clarify the role of the HTH domain of EL222, we measured the TG signal of a short construct that lacks the HTH domain (EL-LOV, residues 1−144) with and without DNA, shown in Figure S2. The two signals are the same, indicating that EL-LOV does not interact with DNA in the dark and light states. Therefore, the interaction site of photoexcited EL222 with DNA is assigned to the HTH domain in the context of the full-length molecule. Dynamics of the Reaction of EL222 with DNA. The reaction scheme for the DNA binding process was established by analyzing the time development of the diffusion signal. Figure 3 shows the diffusion signals observed at various q2

Scheme 2

binding and the rate constant of protein dimerization, respectively. The temporal profiles of the refractive index changes are derived by solving the diffusion-reaction equations (SI-3). We tried to fit the observed diffusion signals on the basis of these schemes. Because there are many parameters in these equations, we had to fix some of them to determine reliable parameters. First, the refractive index and diffusion coefficient of DNA (δnDNA and DDNA, respectively) were reported previously (δnDNA/δnprotein ≈ 1.5, and DDNA = 6.0 × 10−11 m2 s−1 for the 45-mer),17,18 and we used these values. Second, we considered that DR and DI were identical to those of EL222 in the absence of DNA (DR = 8.8 × 10−11 m2 s−1, and DI = 8.5 × 10−11 m2 s−1).13 This assumption is reasonable because DNA does not interact with EL222 in the ground state and the initial reaction is not changed by the presence of the DNA, as shown above. Initially, we fitted the diffusion signals on the basis of Scheme 1. Because in Scheme 1, DNA binding was slower than protein dimerization, it is reasonable to consider that the protein dimerization reaction was not affected by the presence of DNA. Therefore, we fixed δnR, δnI, δnP, DR, DI, DD, and kdimer to the values obtained from the analysis of the dimerization reaction of EL222 without DNA. However, the TG signals could not be reproduced by the model (Figure S3). Next, we fitted the diffusion signals on the basis of Scheme 2. In this analysis, we assumed that kdimer was the same as that previously determined without DNA (kdimer = 8.6 × 103 M−1 s−1). We show below that this estimation is reasonable. Under these restrictions, the signals were successfully reproduced and the following diffusion coefficients and kDNA were determined: DR = (8.8 ± 0.2) × 10−11 m2 s−1, DI = (8.5 ± 0.3) × 10−11 m2 s−1, DI:DNA = (6.5 ± 0.4) × 10−11 m2 s−1, DP = (5.7 ± 0.3) × 10−11 m2 s−1, and kDNA = 4.6 ± 0.5 s−1 (200 μM DNA) (Figure 3). In addition to the quantitative analysis described above, one may qualitatively understand that Scheme 2 is appropriate for describing the reaction by comparing the diffusion signals of EL222 with DNA to those of EL222 without DNA (Figure 3 and Figure S4). In the absence of DNA, the diffusion peak is very weak around 100 ms (Figure S4). This observation indicates that dimerization is negligible in this time range. On the other hand, the diffusion signal clearly appeared in this time range for the EL222 sample with the DNA (Figure 3). Therefore, these observations clearly showed that DNA binding occurs before protein dimerization. Previously, the co-crystal structure of DNA bound to a monomeric fork head domain, a relative of HTH domains with substantial loop extensions, has been reported.19 According to this report, we consider that the temporal DNA binding with the monomer HTH domain is possible. We analyzed the observed TG signals at various EL222 concentrations based on Scheme 2 (Figure 2a). The signals were fitted by eq S2 with a free parameter of kdimer, and the bimolecular reaction rate constant of protein dimerization was determined to be (9.3 ± 0.5) × 103 M−1 s−1 (Figure 2b). This is almost the same as k′dimer, which was measured in the absence of DNA (8.6 × 103 M−1 s−1). The previous assumption that the

Figure 3. TG signals of EL222 with DNA at various q2 values. The concentrations of EL222 and DNA (sequence 1) are 200 μM. q2 = 8.0 × 1012, 2.9 × 1012, 1.1 × 1012, 2.8 × 1011, 9.5 × 1010, 7.0 × 1010, and 4.2 × 1010 m−2 from left to right, respectively. Curves fit by eq S2 are shown as dashed lines.

values. The signals were again normalized by the intensity before the diffusion component (∼0.1 ms). The diffusion signal intensity became stronger with an increasing observation time range (decreasing q2). This q2 dependence is a clear indication that the D change occurs in the observation time window. For this measurement, we used a sufficiently low light intensity for the excitation and the fraction of the excited EL222 was much smaller than the concentration of DNA to ensure that dimerization occurred between the monomers in the light and dark states. We may consider two possible reaction schemes for producing the (EL222*−EL222)DNA complex: dimerization occurring before DNA binding (Scheme 1) and DNA binding occurring before dimerization (Scheme 2). In Schemes 1 and 2, R, I, D, I:DNA, and D:DNA denote the reactant, the intermediate (initial photoexcited EL222), the photoproduct (dimer) of EL222, a complex between EL222* and DNA, and the final photoproduct, respectively. Rate constants kDNA and kdimer represent the rate constant of DNA Scheme 1

D

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry presence of DNA does not change the rate of the dimerization process is therefore appropriate. The reduction of the diffusion coefficient caused by forming I:DNA (DR/DI:DNA = 1.35) may be explained by the increase in molecular size and the intramolecular conformational change. According to the Stokes−Einstein relationship, the ratio of the diffusion coefficient is given by the cubic root of the ratio of the molecular weight. In this case, the DR/DI:DNA ratio is slightly larger than {[MWEL222(24 kDa) + MWDNA(28 kDa)]/ MWEL222(24 kDa)}1/3 = 1.29. The decrease in the diffusion coefficient is therefore due to not only the size increase but also a possible intramolecular conformational change, which increases the friction for the diffusion. The observed large D change is consistent with the previously suggested conformational change of EL222 associated with DNA. On the basis of the crystal structure in the dark, it has been proposed that EL222 will have an open conformation when it binds DNA, while it has a closed conformation in the dark.4 Previously, we found that the yield of EL222 dimerization is low in the absence of DNA.13 This low yield was explained in terms of inhomogeneous conformations of EL222. There are two conformations in the photoexcited state; one can produce the dimer (reactive), and the other cannot (nonreactive). The low dimerization yield can be explained by there being a small fraction of the reactive light state. It could be possible that the reactivities are different for the DNA binding reaction; we therefore examined this possibility. First, it should be noted that the diffusion signals were reproduced by eq S2, which does not take into account the reactive and nonreactive species. If some of the photoexcited EL222 molecules do not bind with the DNA, the diffusion of free EL222* should be observed in the signal. However, no appropriate component was observed. Furthermore, as shown in Figure 1, the intensity of the molecular diffusion signal is significantly increased by the presence of DNA. Because the intensity of the molecular diffusion signal reflects the yield of reactions, the strong peak intensity suggests a high yield of DNA binding. DNA Binding Rate. To determine the second-order rate constant of both DNA binding and dimerization, we measured the TG signals at various concentrations of DNA ([DNA]) at a constant protein concentration ([EL222]). Figure 4a shows the TG signals of EL222 at 50 μM with 10−50 μM DNA. The signals were normalized by the amount of photoactivated molecules. It can be seen that the shape and intensity of the diffusion signal strongly depend on the DNA concentration. The higher the concentration, the stronger the signal. This indicates that the rate of reaction of the DNA binding increases with an increasing concentration. If the DNA binding occurs via a pseudo-first-order reaction, the TG signal was represented by eq S2 with kDNA = k′DNA[DNA], where k′DNA is the intrinsic bimolecular reaction rate constant of the DNA binding step. These signals were reproduced well by analysis using eq S2 with kDNA as the only adjustable parameter. The values of kDNA are plotted versus DNA concentration in Figure 4b. The linear dependence of kDNA on [DNA] confirms that the initial D change represents the DNA binding reaction. The bimolecular reaction rate constant was determined to be (2.2 ± 0.2) × 104 M−1 s−1 from the slope of this plot. DNA Sequence Dependence of DNA Binding. To establish a detailed mechanism for DNA binding, we investigated the DNA sequence dependence of the DNA binding rate. The three sequences of the DNA fragment shown in Materials and Methods were used. Sequences 2 and 3 have

Figure 4. (a) DNA concentration dependence of the TG signals at a q2 of 5.1 × 1010 m−2. The concentrations of DNA (sequence 1) were 10, 20, 30, 40, and 50 μM, with a constant EL222 concentration (50 μM). The best fit curves using eq S2 are shown as dashed lines. (b) Plot of the reaction rate of the DNA binding step vs DNA concentration. The data are fitted with a linear function.

the highest and lowest (no specificity) affinities, respectively. The lengths of the DNA fragments were the same (45 bp). First, the affinity of the protein−DNA interaction was measured using the EMSA method at 23 °C (Figure S5 and Figure 5). As reported previously, EL222−DNA (sequence 1)

Figure 5. Binding affinity measured by an EMSA for sequences 1−3 in the light and sequence 1 in the dark at 23 °C and fractions of bound DNA plotted vs EL222 concentration. Red squares, blue circles, and green triangles represent data for sequences 1−3, respectively. The best fit curves from a Hill equation are shown as solid lines.

interaction was negligible in the dark at all concentrations, while EL222 bound to DNA under light illumination.4 All of the examined fragments of DNA bound to EL222 at high concentrations in the illuminated state. The fraction of the bound state for each sequence is plotted versus protein concentration in Figure 5. The concentrations of EL222 at halfmaximum binding (EC50) were determined to be 13 μM for sequence 1, 2.4 μM for sequence 2, and 18 μM for sequence 3. This order is consistent with the previous reports.4,6 E

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry The kinetics of the DNA binding process were measured from the DNA concentration dependence of the TG signals for each sequence. The signals at 50 μM EL222 and 10−50 μM DNA are shown in Figure 6a. Although the affinities of the

Figure 7. Kinetics of dark recovery monitored by the absorption at 450 nm. The concentrations of (a) EL222 and (b) EL-LOV are 100 μM, and the concentration of DNA (sequence 2) is 100 μM. In panel b, the two signals almost overlap. The light state was established by blue light excitation for 1 min from an LED light source. The best fit curves from a biexponential function are shown as dashed black lines.

the dark recovery kinetics of EL222 are expressed by a biexponential function.5 The dark recovery kinetics with DNA are also reproduced by a biexponential function. The observed lifetimes of EL222 without DNA are 27 and 10 s and with DNA are 230 and 29 s. The lifetimes and amplitudes are listed in Table 1.

Figure 6. (a) DNA sequence dependence of the TG signals at a q2 of 5.1 × 1010 m−2. Red, blue, and green signals represent data for sequences 1−3, respectively. For each sequence, the concentrations of DNA fragments were 10, 20, 30, 40, and 50 μM, with a constant EL222 concentration (50 μM). (b) Plot of the reaction rates of the DNA binding step vs DNA concentration. The data are fitted with a linear function.

Table 1. Kinetics of Dark Recovery Monitored by the Change in Absorptiona

DNA sequences are different, the concentration dependences of the signals were very similar to each other. Using the analyses described above, the intrinsic bimolecular reaction rates of DNA binding were determined for each sequence (Figure 6b): (2.2 ± 0.2) × 104 M−1 s−1 for sequence 1, (2.3 ± 0.2) × 104 M−1 s−1 for sequence 2, and (2.1 ± 0.2) × 104 M−1 s−1 for sequence 3. It is interesting that the reaction rate constants of the DNA binding processes are very similar, despite their different DNA binding affinities. Rate of Dark Recovery. EL222 is a photocyclic DNAbinding protein.4 The dark recovery rate may be important for determining the sensitivity of this function.20,21 During the course of this study of EL222−DNA binding, it was found that the lifetime of the photoexcited EL222 (EL222*) state is prolonged by the presence of DNA (sequence 2). Figure 7a shows the kinetics of the dark recovery of EL222 (100 μM) monitored by the absorption change at 450 nm with and without DNA (100 μM, sequence 2). It is apparent that the lifetime of the photoactivated state becomes much longer in the presence of the DNA. For the purpose of comparison, the dark recovery kinetics of the EL-LOV (100 μM) with (100 μM, sequence 2) and without DNA are also shown in Figure 7b. The lifetime of the photoactivated state of EL-LOV does not depend on the presence of the DNA, indicating that increasing the lifetime of the active state is achieved by the interaction between DNA and the HTH domain. As previously reported,

EL222 EL222 with DNA EL-LOV EL-LOV with DNA

τslow (s)

Aslow

τfast (s)

Afast

27 230 42 44

−0.140 −0.036 −0.029 −0.029

10 29 12 12

−0.076 −0.184 −0.178 −0.181

Signals were fitted by a biexponential function. The time constants and the amplitudes of the slow and fast components are listed.

a

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DISCUSSION In this work, we demonstrated the time-resolved detection of the protein−DNA binding process of EL222 by dynamic diffusion detection. So far, kinetic data for protein−DNA binding have been reported on the basis of SPR and timeresolved fluorescence measurements.3,22−24 Although the TG measurements for this system require approximately a few micromolar, these methods are more sensitive and may be able to use more dilute solutions. However, both have associated drawbacks. For example, the SPR method requires that the sample be fixed onto a metal surface, which can lead to artifacts in the data. Furthermore, it is difficult to detect fast kinetics. For fluorescence measurement, labeling with a fluorophore is required, which may change the kinetics. On this point, the TG method does not require sample fixing or chemical labeling, F

DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry

Although the yield of the dimerization of EL222 without DNA is low, and this low yield was explained in terms of inhomogeneous conformations of EL222, we found that the yield of EL222 dimerization after binding to DNA is high. This suggests that the initial DNA binding step enhanced protein dimerization. When the putative DNA-binding helices (2α,3αhelix) on the HTH domain interact with DNA, the LOV domain is relocated and the whole structure of the protein may change to give an open conformation.4 This open conformation could be favorable for dimerization. We also found that the yield of the DNA binding step is high, in spite of the low yield of the dimerization without DNA. The structure of the EL222* monomer can therefore interact with DNA, although it cannot be dimerized. According to the dark structure, the putative DNA interaction region (2α,3α-helix on the HTH domain) is located on the outer face, while the putative dimerization sites (β-sheet on the LOV and 4α-helix on the HTH domain) are completely buried in the inner part of the protein.4 A small conformational change upon light excitation is enough to bind DNA; however, it is insufficient to form a dimer with a ground state EL222 monomer. Further conformational change (opening of the protein), associated with DNA binding, occurs to make dimerization favorable. Finally, we found that DNA binding stabilizes the active state of EL222 to extend the lifetime. Previous reports demonstrated that the lifetime of the activated state is related to the light sensitivity of proteins; the longer the lifetime, the higher the sensitivity.20,21 At the same time, the short lifetime has an advantage in terms of rapid dark adaptation in a changing environment. In the case of EL222, the longer lifetime of the DNA-binding state enables more efficient transcription, while the fast recovery in the DNA-free state may facilitate rapid dark adaptation to prepare for the next stimulation.

and consequently, reactions of biomolecules can be monitored repeatedly under natural conditions.3,16 Many DNA-binding proteins achieve binding by forming dimers. For example, TraR contains a HTH domain, GCN4bZIP contains a basic region leucine zipper (bZIP) domain, and Gal4 contains a Zn2Cys6 binuclear cluster DNA recognition element, all of which bind with DNA through a dimer.25−27 However, there are some reports on the kinetics of DNA binding.24,28,29 For example, Fos and Jun DNA-binding proteins containing a bZIP element bind to DNA with a heterodimer to mediate activated transcription. Stopped-flow fluorescence resonance energy transfer (FRET) revealed that the two protein monomers bind DNA and subsequently dimerize upon DNA binding.24 In contrast, some proteins initially dimerize and subsequently bind to DNA.16,30 For example, it was found by using the time-resolved diffusion method that the monomeric Aureochrome-1 (AureoCS) from Vaucheria f rigida, which consists of the LOV domain and the DNA-binding bZIP domain, leads to dimerization and subsequently binds with DNA upon photoexcitation.16 The observed DNA-binding-then-dimerization reaction for EL222 indicates that a surface of the HTH domain, which was masked by the LOV domain in the dark state,4 is exposed for DNA binding as a monomer, and subsequent dimerization is important for stabilization of the EL222−DNA complex. The mechanism described above may be related to the observed association and dissociation rates. Interestingly, the reaction rate constant of the DNA binding step was independent of the DNA sequence, regardless of binding affinity. Because the binding affinity is determined by a ratio of the rate constants of DNA binding (kon) and dissociation (koff), this indicates that the specificity of DNA binding is mainly regulated by the dissociation rate. The sequence-independent binding rate suggests that the binding process is controlled by protein−DNA collision and electrostatic interaction. Once the complex is formed, the attractive interaction stabilizes the bound state if the DNA has the target sequence. However, if the DNA does not have the target sequence (nonspecific binding), the attractive interaction is weak and results in faster dissociation. Dimerization after DNA binding may be important for this stabilization. This mechanism is reasonable for DNA recognition. A similar type of DNA recognition was suggested for the DNA binding/dissociation of the c-Myb DNA-binding domain (R2R3*) investigated using SPR. Because the rate of DNA binding (kon) was not affected by the DNA sequences despite their different DNA binding affinities,22,31 it was proposed that c-Myb nonspecifically binds to the DNA first. If the DNA sequence is the target one, specific hydrogen bonds and van der Waals interactions lead to a slow dissociation rate. On the other hand, a different mechanism was reported for the zinc finger DNA transcription factor (ADR1 PAR-ZF), which has a very similar protein−DNA complex lifetime regardless of the DNA sequence. This suggests that the binding specificity comes mainly from the difference in the association phase.23 The kinetic values for DNA binding for these proteins showed no clear differences [2.1−2.3 × 104 M−1 s−1 for EL222 (DNA fragments were 45-mers), 2.1−2.5 × 105 M−1 s−1 for c-Myb R2R3* (22-mer), and 5.6 × 103 to 1.9 × 105 M−1 s−1 for ADR1 PAR-ZF (26-mer)].22,23 At present, it is not clear what factors cause these differences in the DNA binding scheme; however, it will be important to focus future work on the molecular origin of DNA binding.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.7b01206. Laser power dependence of the TG signal (SI-1), TG signal of EL-LOV with DNA (SI-2), analytical models (SI-3 and SI-4), the TG signal of EL-LOV (SI-5), and the EMSA measurements (SI-6) (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Masahide Terazima: 0000-0001-6828-479X Funding

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

The authors declare no competing financial interest.

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REFERENCES

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DOI: 10.1021/acs.biochem.7b01206 Biochemistry XXXX, XXX, XXX−XXX