Hydrogen Bonding Environment of the N3âH Group of Flavin

Article

Hydrogen Bonding Environment of the N3–H Group of FMN in the LOV domains of Phototropins Tatsuya Iwata, Dai Nozaki, Atsushi Yamamoto, Takayuki Koyama, Yasuzo Nishina, Kiyoshi Shiga, Satoru Tokutomi, Masashi Unno, and Hideki Kandori Biochemistry, Just Accepted Manuscript • Publication Date (Web): 22 May 2017 Downloaded from http://pubs.acs.org on May 27, 2017

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Biochemistry

Hydrogen Bonding Environment of the N3–H Group of FMN in the LOV Domains of Phototropins Tatsuya Iwata,†,# Dai Nozaki,† Atsushi Yamamoto,† Takayuki Koyama,† Yasuzo Nishina,‡ Kiyoshi Shiga,§ Satoru Tokutomi,|| Masashi Unno,⊥ and Hideki Kandori*,† †

Department of Life Science and Applied Chemistry, Nagoya Institute of Technology, Showaku, Nagoya 466-8555, Japan ‡

Department of Molecular Physiology, Graduate School of Medical Sciences, Kumamoto University, Honjo, Kumamoto 860-8556, Japan

§

Department of Physiology, School of Health Sciences, Kumamoto University, Kuhonji, Kumamoto 862-0976, Japan

||

Department of Biological Science, Graduate School of Science, Osaka Prefecture University, Sakai, Osaka 599-8531, Japan ⊥

Department of Chemistry and Applied Chemistry, Graduate School of Science and Engineering, Saga University, Saga 840-8502, Japan

ACS Paragon Plus Environment

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Abstract The light-oxygen-voltage (LOV) domain is a flavin-binding blue-light receptor domain, originally found in a plant photoreceptor phototropin (phot). Recently, LOV domains have been used in optogenetics as the photosensory domain of fusion proteins. Therefore, it is important to understand how LOV domains exhibit light-induced structural changes for the kinase domain regulation, which enables the design of LOV-containing optogenetics tools with higher photoactivation efficiency. In this study, the hydrogen bonding environment of the N3–H group of FMN of LOV2 domain from Adiantum neochrome (neo) 1 were investigated by lowtemperature Fourier transform infrared spectroscopy. Using specifically 15N-labeled FMN, [1,315

N2]FMN, the N3–H stretch was identified at 2831 cm–1 for the unphotolyzed state at 150 K,

indicating that the N3–H group forms a fairly strong hydrogen bond. The N3–H stretch showed temperature-dependence, with a shift to lower frequencies at ≤200 K and to higher frequencies at ≥250 K from the unphotolyzed to the intermediate states. Similar trends were observed in the LOV2 domains from Arabidopsis phot1 and phot2. By contrast, the N3–H stretch of Q1029L mutant of neo1-LOV2 and neo1-LOV1 were not temperature dependent in the intermediate state. These results seemed correlated with our previous finding that the LOV2 domains show the structural changes in the β-sheet region and/or the adjacent Jα helix of LOV2 domain but that such structural changes do not take place in Q1029L mutant or neo1-LOV1 domain. The environment around the N3–H group was also investigated.


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Introduction Flavins are utilized as chromophores in some types of photoreceptors in organisms.1 The lightoxygen-voltage (LOV) domain is a flavin-binding photoreceptive domain, which was originally found in the photoreceptor domains of phototropin (phot),2, 3 a plant blue-light receptor. The LOV domain is named after the homology of the primary2 and tertiary4 structures to bacterial light-sensor photoactive yellow protein,5 oxygen sensor FixL,6 and the N-terminal domain of a voltage-gated potassium channel, HERG,7 the protein fold of which belongs to the Per-Arnt-Sim (PAS)L superfamily.8

Phot is a photoreceptor in plants where it is related to phototropic

response,9 relocation of chloroplasts10,

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and stomata opening,12 and other functions. Phot

comprises about 1000 amino acid residues and two flavin mononucleotide (FMN) molecules, where the two LOV domains and FMN are at the N-terminal side and the Ser/Thr kinase domain is at the C-terminal side2. In addition to phots, LOV domains are found in other blue-light photoreceptors such as FKF113 and aureochrome.14 The LOV domains located at the N-terminal side regulate functional domains at the C-terminus, such as in phots, but vice versa in aureochromes. The two LOV domains have different functions although they have similar amino acid sequences.

The kinase activity is primarily regulated by the photoreaction of the LOV2

domain.15, 16 The LOV1 domain is referred to as the dimerization17, 18 and light attenuation site.16 Therefore, differences in the structure and structural changes in both LOV domains are of interest. The X-ray crystal structures of the LOV1 and LOV2 domains are quite similar, and there is no explanation for why the two LOV domains have different roles based on structural features.4, 19-21 The approximate positions of the LOV2 and kinase domains were determined by


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X-ray small angle scattering22,

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in order to estimate how the LOV domain(s) regulates the

activity of the kinase domain. In 2003, an NMR study demonstrated the existence of an α-helix at the C-terminus of LOV2 and the α-helix regulates kinase activity.24 Structural changes in the α-helix were also studied using other techniques, including transient grating25-28 and Fourier transform infrared (FTIR) spectroscopy.29, 30 Our previous FTIR study showed that the structural changes in the Jα helix (or the corresponding region at the C-terminus of the LOV2 domain) differ among the LOV2 domains in Arabidopsis phot1 and phot2, and Adiantum neochrome 1 (neo1).30 Previous results suggest that the structural changes in Jα are specific to phot1 and phot2. The structural changes may differ between LOV1 and LOV2, and among the LOV2 domains of phot1, phot2, and neo1, but the photochemical reaction is generally the same in the LOV domains, where an adduct forms between FMN and a nearby cysteine.31-34 FMN absorbs light and after a singlet excited state of FMN, intersystem crossing occurs over a nanosecond timescale.35 Next, a shorter wavelength product appears with a time constant of the microsecond timescale,33, 36 which is called S390; this is the adduct formed between FMN and a cysteine. A recent study reported the existence of a pathway from the singlet excited state to the adduct form in the LOV2 domain of Chlamydomonas phot.37 S390 reverts to the unphotolyzed state in the dark within seconds to hours, where the time varies among LOV domains.31,

38, 39

Various

models have been proposed for the reactive cysteine, but previous FTIR studies indicate that the cysteine is protonated in both the unphotolyzed39-42 and the triplet excited43 states of FMN. Dynamic protein motion probably plays an important role in adduct formation.44 Recently, LOV domains have been utilized as photosensory domains in fusion proteins for use in optogenetics or light manipulation.45-47 Optogenetics is a technique for manipulating neurons


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or other cells in organisms by light illumination, where light-gated ion channels (channelrhodopsin) or other light-reactive enzymes are expressed. Therefore, it is important to determine how LOV domains exhibit light-induced structural changes in order to understand the appropriate regulation of these functional domains in optogenetics tools. The light-dependent regulation of a LOV domain as a functional domain in a fusion protein was demonstrated previously based on the α-helical movement of Jα helix in the LOV2 domain.47 However, the different structural changes of LOV2 domains suggest that diverse regulation systems may be employed in various types of enzymes.

Thus, some guiding principles are required for

optogenetics tools when designing fusion proteins with LOV domains and catalytic domains with various structures. In this study, in order to understand how the photochemical reaction of the FMN chromophore induces structural changes in LOV domains, we investigated the hydrogen bonding environments of FMN by FTIR spectroscopy. FMN forms hydrogen bonds with apoprotein via its C2=O, C4=O, and N3–H groups. We focused on the hydrogen bonds of N3–H stretch because we previously determined the hydrogen bonding environments of two C=O stretches in the FMN chromophore of Adiantum neo1-LOV2.48

We identified multiple temperature-dependent

structures for the S390 intermediate using low temperature FTIR spectroscopy in various LOV domains, including mutants.39 15

49-52

Thus, we prepared neo1-LOV2 reconstituted with [1,3-

N2]FMN in order to identify N3–H stretch and to investigate how the N3–H group changes its

hydrogen bonding environment. N3–H stretch was assigned to 2831 (–)/2812 (+) cm–1 measured at 150 K. We also identified the temperature dependence of the structural changes in N3–H stretch and the incapacity for the exchange of hydrogen for deuteron under D2O conditions. Similar changes were observed in Arabidopsis phot1-LOV2 and phot2-LOV2 despite the


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presence of Jα helix. However, these changes in the hydrogen bonding strength were not observed in a Q1029L mutant, dehydrated neo1-LOV2, and neo1-LOV1, which did not exhibit progressive structural changes.39, 50 51 The hydrogen bonding environment of the N3–H group was strongly correlated with conformational changes in the peptide backbone.

Materials and Methods Preparation of Various LOV2 Domains.

Adiantum neo1-LOV2 (Pro905–Pro1087) and its

Q1029L mutant, and neo1-LOV1 (Gly660-Val805) were expressed as a fusion protein with calmodulin-binding peptide at the N-terminal side in Escherichia coli strain BL21(DE3) and purified.48-50

Arabidopsis phot1-LOV2-core (Lys462–Arg586), phot1-LOV2-Jα (Lys462–

Asp617), phot1-LOV2-Jα phot2-LOV2-core (Gln376–Gln500), phot2-LOV2-Jα (Gln376– Asp531), Adiantum neo1-LOV2-core (Arg916–Pro1045), and neo1-LOV2-Jα (Arg916– Asp1076) were expressed as a fusion protein with glutathione S-transferase (GST) at the Nterminal side in BL21(DE3), and the GST tag was removed by digestion with thrombin.29, 30 Specific site-directed 13C- and

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N-labeled FMNs, i.e., [1,3-15N2]FMN, [4,10a-13C2]FMN, and

[2-13C]FMN, were synthesized as described previously.53 Reconstitution of FMN and apoprotein was performed as described previously.48 FTIR Measurements. FTIR measurements were obtained as described previously.48, 49 Infrared spectra of the hydrated films were measured using an FTS-7000 (DIGILAB) spectrophotometer. Hydrated films were illuminated with light at >400 nm using a combination of a halogentungsten lamp (1 kW) and a long-pass filter (L42, Asahi Techno Glass). The accumulated intermediate was only one kind, S390 intermediate, measured by UV-vis spectroscopy.40,

49

Spectral intensity was normalized by use of vibrational bands at 1500–1200 cm–1.54


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Density Functional Theory (DFT) Calculations. The frequencies of N3–H stretch in 7,8dimethyl-10-glycerylisoalloxazine and its C4a–thiol adduct form with and without hydrogen bonding partners were determined from previous DFT calculations.54

The harmonic and

anharmonic vibrational frequencies were calculated using the Gaussian09 program.55

Results 15

N Isotope Effect of FMN on Vibrations at 1800–950 cm−1. In order to identify the vibration

signals where the N3 atom of FMN was involved, we prepared reconstituted neo1-LO2 with [1,3-15N2]FMN. Figure 1 shows the difference FTIR spectra for neo1-LOV2 reconstituted with unlabeled FMN (solid line) and [1,3-15N2]FMN (dotted line) measured at 150 (a) and 295 K (b) in the 1800–950 cm−1 region. At both temperatures, the identical bands exhibited downshifts of 4–10 cm−1, i.e., negative bands at 1507, 1407, 1272, and 1250 cm−1, and positive bands at 1541, 1521, 1379, 1258, and 1197 cm−1 at 150 K. Previously, we attributed the bands at 1551 (−)/1541 (+) and 1521 (+)/1507 (−) cm−1 to C10a=N1 stretching modes.48 However, the negative band at 1551 cm–1 did not exhibit isotope shift. According to the DFT calculations for 7,8-dimethyl-10-glycerylisoalloxazine,54 C10a=N1 stretch in the unphotolyzed and intermediate states were assigned to 1568/1512 and 1542 cm−1, respectively, and the band at 1551 cm−1 was assigned to C=C stretching involving ring I and C8a (see the inset in Figure 1 for details of the atom and ring numbering).


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1541

Difference Absorbance (0.01 abs./div.)

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1521

1507

1379

1258

1197

(a) 150 K

1407 1272

1250

1551 1538 1519

1380 1195

1261

1504

1405

(b) 295 K

1272 1249

1553

1800

1600

1400

1200

1000

-1

Wavenumber (cm ) Figure 1. (a) Difference FTIR spectra of unlabeled neo1-LOV2 (solid line) and uniformly

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N-labeled

(15N-FMN/15N-apoprotein) neo1-LOV2 (dotted line) in the 1800–950 cm−1 region measured at 150 K. (b) Difference FTIR spectra of neo1-LOV2 reconstituted with unlabeled FMN (solid line) and 15N-labeled (dotted line) apoprotein in the 1800–950 cm−1 region at 150 K. The inset shows the positions of the labeled nitrogen atoms in the isoalloxazine ring.


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The negative bands at 1407, 1272, and 1250 cm−1 exhibited isotope shifts of 4–7 cm−1. The lower frequencies suggest that these vibrations contained the single band characteristics of FMN, such as C2–N1, C2–N3, and C4–N3 stretching, and N3–H bending modes. The positive bands at 1379, 1258, and 1197 cm−1 exhibited isotope shifts of 5–9 cm−1. These bands are mainly attributable to ring stretching vibrations due to displacement of the N1 and N3 atoms. The bands at 1379 (+), 1272 (−), 1258 (+), and 1250 (−) exhibited spectral downshift for [4,10a-13C2]FMN and [2-13C]FMN,48 so they may be attributable to C4–N3, C2–N1, and C2–N3 stretches. According to DFT calculations by Kikuchi et al., these bands originate from C–N stretches in rings II and III of the isoalloxazine ring.54

The N3–H bending modes were calculated at

1443/1436 and 1445/1441 cm−1 in the presence of a hydrogen bonding acceptor for the unphotolyzed and S390 intermediate states, respectively. It is likely that the small frequency shift of the N3–H bending mode between the unphotolyzed and intermediate states prevented us from detecting the FTIR bands of N3–H bending in the difference FTIR spectra.

Identification of the N3–H Stretching Vibrational Mode. The higher frequency region was investigated to identify N3–H stretch in the FTIR spectra.

N3–H stretch of riboflavin

tetraacetate appeared at 3380 and 3200 cm−1 in chloroform solution, which are the hydrogen bonding-free and hydrogen-bonded forms, respectively.56 However, we found N3–H stretch in a much lower frequency region than 3200 cm−1. Figure 2 shows the difference FTIR spectra in the 3000–2740 cm−1 region where C–H stretches mainly appear. The peaks are as follows: 2970 (+), 2964 (–), 2949 (+), 2936 (−), 2929 (+), 2905 (+), 2878 (−), 2873 (+), 2864 (+), 2857 (−), 2831 (−), and 2812 (+) cm−1 (dotted lines in Figure 2a–e). It should be noted that both spectra agreed very well with each other for the unlabeled FMN-reconstituted and unreconstituted (native


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formation of holoprotein in bacteria) LOV2 domains in this frequency region (Figure S1). In this region, the band at 2831 (−)/ 2811 (+) cm−1 exhibited downshift to 2816 (−)/2799 (+) cm−1 for [1,3-15N2]FMN-reconstituted noe1-LOV2 (Figure 2a). Bands appeared at 2816 (−)/2799 (+) cm−1 and at 2831 (−)/2811 (+) cm−1 for the uniformly 15N-labeled neo1-LOV2 (Figure 2b) and the reconstituted neo1-LOV2 of

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N-FMN/15N-apoprotein (Figure 2c), respectively.

These

results indicate that the bands originate from the N3–H stretch of FMN. We could not exclude the possibility that a C–H group was influenced by 15N-labeling because C–H stretching modes appear in this region. Figure 2d and e show the difference spectra for 13Clabeled neo1-LOV2 and 12C-FMN/13C-apoprotein, respectively. The negative band at 2831 cm−1 exhibited downshift of 2 cm−1 for the

13

C-labeled neo1-LOV2 (Figure 2d), while a band

appeared at 2831 cm−1 for the 12C-FMN/13C-apoprotein (Figure 2e). The positive band at 2811 cm−1 did not exhibit downshift. The downshift of that of

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C-labeled neo1-LOV2 was different from

N-labeled neo1-LOV2, so the bands at 2831 (−)/2811 (+) cm−1 originated from N–H

stretch rather than C–H stretch. The downshift of the N3–H stretching vibrational bands by 13C-labeling was affected indirectly by the

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C-labeling of FMN.

In fact, the band downshifted by 2–3 cm−1 for neo1-LOV2

reconstituted with [4,10a-13C2]FMN and [2-13C2]FMN (Figure 2f and g, Table 1). These results show that 13C-labeling of the atoms neighboring FMN influenced N3–H stretch. The other bands in the 2970–2857 cm−1 region were downshifted by 4–10 cm−1 for the uniformly

13

C-labeled

neo1-LOV2, thereby indicating that these bands originated from C–H stretching modes (Figure 2d). These bands also downshifted for the

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C-FMN/13C-apoprotein (Figure 2e), so they were

due to apoprotein and the C–H groups from the amino acid side chains.


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Figure 2. Difference FTIR spectra for unlabeled (dotted lines in a, c, e–g), native (broken lines in b, and d), and labeled (solid lines) neo1-LOV2 in the 3000–2740 cm−1 region measured at 150 K. (a) [1,315

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N2]FMN reconstituted, (b) uniformly

C-labeled, (e)

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15

N-labeled, (c)

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N-FMN/15N-apoprotein, (d) uniformly

C-FMN/13C-apoprotein, (f) [4,10a-13C2]FMN reconstituted, and (g) [2-

13

C]FMN reconstituted neo1-LOV2 are shown.


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Table 1. Observed and calculated N3–H stretch in the unphotolyzed state.

Experimental

a, b

DFT calculationc, d model 1

model 2

2831 (–15, –3, –2) (150 K)

harmonic approximation

3473 (–8, 0, 0)

3114 (–8, 0, 0)

2821 (–18, –5, –7) (295 K)

anharmonic calculation

3447

2972 (–6, –2, 3)

model 3

model 4f

intermediate state

Experimental

a, b

DFT calculationc, d

3183 (–2, 0, 0) harmonic approximation

3477 (–8, 0, 0)

2812 (–13, –2, n.d.e)

3138 (–6, 0, 0)

2839 (–7, –5, –5)

3081 (–9, 1, –5) anharmonic calculation

3352 3035 (–8, –5, –7)

a

The numbers in parentheses are the isotope shifts for [1,3-15N2]FMN, [4,10a-13C2]FMN, and

[2-13C]FMN, respectively. bThe top and bottom frequencies are those measured at 150 and 295 K, respectively. cThe numbers in parentheses are the isotope shifts for [1,3-15N2]-na, [4,10a13

C2]-na, and [2-13C]-na, respectively. dThe model structures for the DFT calculations were

obtained from Kikuchi et al.54

DFT Calculation of N3–H Stretch in the Absence and Presence of Hydrogen Bonding Partners. DFT calculations were performed to investigate the effects of hydrogen bonding around the N3– H group of FMN. According to previous DFT calculations for N3–H stretch in 7,8-dimethyl-10glycerylisoalloxazine, N–H stretch was identified at 3473 and 3115 cm−1 in the absence and presence of a hydrogen bonding partner, respectively, i.e., the C=O group of Asn998 (models 1 and 2 given by Kikuchi et al.,54 Table 1). This suggests the existence of a strong hydrogen bond


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at the N3–H group due to the presence of hydrogen bonding acceptor(s) because the N–H and O– H stretch in the lower frequency region indicates the formation of hydrogen bonds. In addition, it has been reported that large anharmonic effects appear in X–H (X = O, N, C) stretches,57 thereby indicating that the observed lower frequency of N3–H stretch may have originated from anharmonic effects rather than hydrogen bonding effects. Therefore, a DFT calculation that considered the anharmonicity was performed (Table 1). The anharmonic frequency of N3–H stretch for 7,8-dimethyl-10-glycerylisoalloxazine was calculated at 3447 and 2972 cm−1 in the absence and presence of hydrogen bonding acceptors, respectively. A lower frequency shift was also shown by the harmonic calculation. The differences between the harmonic and anharmonic frequencies were 26 and 143 cm−1 in the absence and presence of hydrogen bonding acceptors, respectively. The frequency at 2972 cm−1 is much closer to the observed value (2831 cm−1). This suggests that the lower frequency shift of N3–H stretch also originated from the increased anharmonicity as well as from hydrogen bonding formation. In the adduct form of the model compound (model 3 given by Kikuchi et al.54), N3−H stretch appeared at 3477 and 3352 cm−1 without the hydrogen bonding partners based on calculations of the harmonic approximation and anharmonic contribution, respectively. In the adduct form, the anharmonic effect might not be negligible because the frequency difference at 125 cm−1 was larger than the frequency difference for the non-adduct form. With a hydrogen bonding donor, the N3–H group exhibited vibrational coupling with the N–H group from Asn980, thereby resulting in two frequencies of N3–H stretch, i.e., 3183 and 3139 cm−1, and 3081 and 3035 cm−1. Thus, the N3–H stretch appeared in a region at a frequency that was ~300 cm−1 lower in the presence of the hydrogen bonding acceptors.


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The isotope effect of N3–H stretch due to

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C labeling of nearby carbon atoms in FMN was

identified by the anharmonic calculation but not by the harmonic calculation (Table 1). The trend in the calculated frequency shift was similar to that measured, which indicates that the anharmonicity is not negligible.

H/D Exchange of the N3–H Group. Figure 3 shows the difference FTIR spectra for hydrated films prepared with H2O (solid lines) and D2O (dotted lines) in the 3000–2740 (a) and 1800–950 (b) cm−1 regions at 150 K. When the N–H group was deuterated, the band at 2831 cm−1 was expected to appear at ~2070 cm−1. However, the band at 2831 (−)/2812 (+) cm−1 remained in these conditions (Figure 3a) and no band was observed in the 2250–2000 cm–1 region (see Figure 2 in Nozaki et al.58). Thus, the N3–H group was not deuterated by the hydration of D2O in neo1LOV2. In our experimental conditions, H/D exchange was moderate so the unchanged X–H groups formed strong hydrogen bonds in proteins. As shown above, it is reasonable to assume that N3–H formed a very strong hydrogen bond so H/D exchange did not occur. In the 1800–950 cm−1 region, an H/D exchange effect was observed in the 1660–1620 and 1300–1250 cm−1 regions and the 1084 (−) cm−1 band. The N3–H group was not deuterated, so this effect originated from the deuteration of the exchangeable X–H groups in the peptide backbone (N–H groups) and of the side chains (N–H and O–H groups). The decreases in the intensities of the bands at 1660 (+), 1645 (−), and 1627 (+) cm−1, and the appearance of the band at 1639 (+) cm−1 are attributable to the amide-I vibrational mode, which was affected by the N–D groups of the peptide backbone. The spectral differences at 1300–1250 cm−1 are attributable to the amide-III vibrational mode.


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Figure 3. Difference FTIR spectra for the unlabeled neo1-LOV2 hydrated with H2O (solid lines) and D2O (dotted lines) in the 3000–2740 cm−1 region (a) and 1800–950 cm−1 region (b) measured at 150 K.

Swartz et al. showed that there was an H/D effect according to the difference FTIR spectra obtained for oat phot1-LOV2 in the 1300–1200 cm−1 region.59 They suggested that the N3–H group is accessible to bulk protons via protein residues or structural water molecules based on their experimental pH-dependent titration of fluorescence.33 In the case of neo-LOV2, the shifts were not due to deuteration of the N3–H group, although structural waters were exchanged for D2O.58 It is unlikely that H/D exchange can occur readily for the N3–H group in LOV domains. Ataka et al. suggested the possibility of asymmetric P–O stretches in the phosphate group of


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FMN based on their study of Chlamydomonas phot-LOV1.41 The origin of the negative band at 1084 cm–1 is unclear because it disappeared or shifted elsewhere.

Temperature Dependence of the Vibrations in the 3000–2740 cm−1 Region. Difference FTIR spectra were measured at 200, 250, and 295 K in addition to 150 K (Figure 4), where different structural changes in the peptide backbone were observed despite the same chromophore.48, 49 As shown in Figure 4a (reproduced from Figure 2a), at 150 K, the band for N3–H stretch appeared at 2831 (−)/2812 (+) cm−1. A similar signal was also observed at 2828 (−)/2805 (+) cm−1 at 200 K (Figure 4b). N3–H stretch of the S390 intermediate state appeared at lower frequencies compared with that of the unphotolyzed state. Moreover, N3–H stretch shifted to higher frequencies after the formation of S390 at 250 and 295 K, i.e., to 2843 (+)/2824 (−) and 2839 (+)/2821 (–) cm−1, respectively (Figure 4c and d). The hydrogen bonds of the N3–H group in the S390 state were weaker at ≥250 K than at ≤200 K. The structural change in FMN (FMN-cysteinyl adduct) did not occur between 200 and 250 K for the S390 intermediate, so the change in the N3–H group was caused by apoprotein rather than by FMN itself. The hydrogen bonding environment of the N3–H group for the S390 state was temperature dependent. The C=O group of Asn998 (for Adiantum neo1-LOV2) is a partner in the hydrogen bond of the N3–H group, so the environment around Asn998 must be changed. Asn998 comprises a β-strand and the transition temperature of the structural changes in the N3– H group corresponded to that at which the structural changes in the β-sheet region appeared. Therefore, the hydrogen bonding strength of the N3–H group reflected the structural change in Asn998.


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A temperature-dependent effect was also observed in the 3000–2850 cm–1 region, which originated from apoprotein (see Figure 2d and e). The spectrum obtained at 200 K was similar to that at 150 K, but the spectra were different at 250 and 295 K. The bands at 2979 (+), 2958 (–), 2949 (+), 2920 (–) 2876 (+), and 2860 (–) cm−1 at 295 K were also assigned to the C–H stretching vibrations of apoprotein based on the uniformly reconstituted neo1-LOV2 of

12

C-FMN/13C-apoprotein.

13

C-labeled neo1-LOV2 and

We determined the temperature

dependence of the amide-I region, which exhibited progressive structural changes in the peptide backbone and the C–H stretching region also reflected the differences in the structural changes with temperature.


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Figure 4. Difference FTIR spectra for neo1-LOV2 reconstituted with unlabeled (solid lines) and [1,3-15N2] (dotted lines) FMN in the 3000–2740 cm–1 region measured at 150 (a), 200 (b), 250 (c), and 295 K (d).


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Correlation between the Hydrogen Bond of N3–H and Structural Changes in the β-sheet in the S390 Intermediate. Previously, we reported that the unhydrated wild type (WT) neo1-LOV2 and Q1029L mutant did not exhibit structural changes in the β-sheet region after the formation of S390 at 295 K, whose structural changes were similar as those of WT neol-LOV2 at lower temperatures.50, 51 Thus, we expected that N3–H stretch would appear at a lower frequency in S390 for these samples as is the case for the measurement of WT at ≤200 K (see Figure 4a and b). Figure 5 shows the difference FTIR spectra for the unhydrated WT (a) and hydrated Q1029L mutant (b) reconstituted with unlabeled (solid lines) and [1,3-15N2]FMN (dotted lines). In this mutant, at 295 K, N3–H stretch appeared in the 2824(–)/2795 (+) and 2834(–)/2814 (+) cm−1 regions for the unhydrated WT and hydrated Q1029L, respectively. The N3–H stretching mode appeared at a lower frequency in S390, which was similar to that determined for the hydrated WT measured at 150 and 200 K (see Figures 2 and 3). These results indicate that there was a correlation between the upshift of N3–H stretch and structural changes in the β-sheet region. Similar trends were observed in the FTIR spectra of the LOV2 domains with other amino acid lengths in Adiantum neo1-LOV2, and in Arabidopsis phot1 and phot2 (Figure S2a–c). Reconstitution of labeled FMN was not conducted, but the bands around 2800 cm–1 were temperature dependent between 150 and 295 K, where the former appeared at lower frequencies and the latter at higher frequencies. By contrast, the N3–H stretch in Adiantum neo1-LOV1 was not temperature dependent (Figure S2d). The hydrogen bonding environment of the N3–H group was very strong among the LOV domains, and the hydrogen bonding strength and structural changes in apoprotein were strongly correlated with each other.


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Figure 5. Difference FTIR spectra for neo1-LOV2 reconstituted with unlabeled (solid lines) and [1,3-15N2] (dotted lines) FMN in the 1800–950 cm−1 region at 150 K (a) and 295 K (b).

Discussion Hydrogen Bonding Environment of the N3–H Group of FMN in Neo1-LOV2. In this study, we assigned the FTIR band for N3–H stretch in FMN and the structural changes after adduct formation in neo1-LOV2. The N3–H stretching band appeared at 2831 and 2821 cm−1 in the unphotolyzed state at 150 and 295 K, respectively. The frequency at which the band appeared


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was quite low for N3–H stretch in flavin compared with that in riboflavin tetraacetate in chloroform at 3380 cm–1.56 The low frequency shift of N3–H stretch was explained by the presence of the hydrogen bonding partner according to the DFT calculations for 7,8-dimethyl10-glycerylisoalloxazine. In fact, the H/D exchange of the N3–H group did not occur under mild deuteration conditions. Previous studies have reported that X–H (X = N, O) stretches appeared at lower frequencies (>3000 cm−1) in some photoreceptive proteins. Thus, the N–H stretch in the retinal protonated Schiff base of bacteriorhodopsin appeared at ~2800 cm−1 and ~3350 cm−1 in the ground state and K intermediate, respectively.60 The positively charged N–H group of the Schiff base interacted with the negatively charged counterion Asp85, so it was expected that the N–H group would form a strong hydrogen bond. The N–H stretch of histidine residue was also observed in the 2800–2600 cm−1 region (as a Fermi resonance signal) for photosystem II61,62 and sensory rhodopsin I.63 In the case of photosystem II, strongly hydrogen bonded histidines were observed sandwiched with Fe2+ and negatively charged quinone,61 or they coordinated with manganese ion in the oxygen evolving complex.62 In the case of sensory rhodopsin I, it is not known whether the positively charged histidine residue had a strong hydrogen bond via its hydrogen bonding partner.63 In both cases, charged species were involved in the strong hydrogen bonds. In BLUF domains, the O–H stretch due to a tyrosine residue appeared at ~2800 cm−1 in the red-shifted intermediate state, where there was quite strong hydrogen bonding, probably with a nearby glutamine residue.64 However, in this case, no charges were apparent in the specific hydrogen bonding environment. At present, it is not clear whether strong hydrogen bonds are formed with the N3–H group in the LOV domain, but this study of BLUF domains might suggest that they are.


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The downshift of the N3–H group due to hydrogen bonding formation was estimated as ~500 cm−1. Dürr et al. reported that riboflavin can bind to oat phot1-LOV2 apoprotein and that the R40D mutant can bind riboflavin.65 Riboflavin does not have a phosphate group and Arg40 is an ionic bond partner of the phosphate group in FMN. Thus, the ionic bond between FMN and apoprotein is not necessarily essential. Our findings suggest that a specific hydrogen bond interaction between the N3–H group of flavin and the C=O group of Asn998 has important roles in the binding of flavin to apoprotein.

Structural Changes in the N3–H Group in the S390 State. The hydrogen bond of the N3–H group was strengthened at ≤200 K but weakened at ≥250 K in the S390 state (Figure 3). Because the accumulated intermediate is only S390, the formation of FMN-thiol adduct, at any temperature measured, this change can be explained not by the structural change of FMN itself but by the interaction between a hydrogen bond partner, which is the C=O group of Asn998.4 This indicates that the structural change in the N3–H group is an indicator of the structural change in Asn998. The simplest interpretation is that the distance between the N3–H group of FMN and the C=O group of Asn998 was closer at ≤200 K but more distant at ≥250 K. Figure 6 shows a schematic model of the hydrogen bonding environment between FMN and apoprotein, which includes our previously reported results for the C2=O and C4=O groups.48 N3–H forms a strong hydrogen bond in the unphotolyzed state (left panel of Figure 6), but the bond becomes much stronger in the S390 intermediate (middle panel of Figure 6). However, the hydrogen bonds weaken for the C2=O and C4=O groups. The crystal structures of the unphotolyzed and S390 states demonstrate that the distance between the oxygen atom of C2=O and the nitrogen atom of the Asn998 side chain increases, whereas the distance between the N3 nitrogen and the


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oxygen atom of the Asn998 side chain decreases.4, 34 In the case of neo1-LOV2, we proposed two states for S390, i.e., S390I and S390II, where S390I exhibits structural change in the turn and α-helix structures, and S390II exhibits structural changes in the β-sheet.66 The crystal structure of S390 probably represents S390I. The resolutions of the crystal structures are 2.7 and 2.6 Å in the unphotolyzed and S390 states, respectively, so the difference of 0.1 Å may not be sufficiently reliable. Our results support the observations based on the crystal structures. In the S390II state, the hydrogen bond of the N3–H group weakens.

The simplest

interpretation of this change is that the distance between Asn998 and FMN increases, and Asn998 probably moves far from FMN. Considering that Asn998 is located in the β-strand and that the structural change occurs in the β-sheet region, then Asn998 moves far from FMN. This strongly suggests that the β-sheet region responsible for the structural change is that containing Asn998. Previously, we showed the importance of Gln1029 for structural changes in the βsheet.50 Gln1029 switches the hydrogen bonding partner after adduct formation, which begins with structural changes in the β-sheet. The structural change of Gln1029, which is located on the βE strand, is then transferred to Asn998 via the βD strand. Phenomena similar to N3–H stretch in FMN have been observed in other LOV2 domains (Figure S2). It is curious that the apparent structural changes in the β-sheet were quite small for the LOV2-core domains in Arabidopsis phot1 compared with the LOV2-core domain in Adiantum neo1. However, the hydrogen bonding environment of the N3–H group is temperature dependent, thereby implying that structural changes occur in the β-sheet or β-strand where the asparagine (Asn544 for Arabidopsis phto1) is located. Thus, common mechanisms may explain the structural changes in LOV2 domains. All LOV2 domains exhibit temperature dependence in terms of the hydrogen bonding strength of the N3–H


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group in the S390 state, which reflects the progressive structural changes in the β-sheet or βstrand where Asp544 is located. However, it is still unclear how the common structural changes in the N3–H group of FMN and the nearby asparagine induce different structural changes, i.e., in the Jα helix for Arabidopsis phto1 and in the β-sheet for neo1-LOV2, and thus the mechanisms responsible must be determined. Elucidating these mechanisms will help to understand the kinase domain activation mechanisms and guide the construction of fusion proteins that can be activated/deactivated by the photoreaction involving LOV domains.

Figure 6. Schematic showing the hydrogen bonding environment between FMN and apoprotein in neo1LOV2 for the unphotolyzed (PDB entry: 1G28) and S390 (PDB entry: 1JNU) states. The peptide backbone is represented as ribbons. FMN is shown as a stick drawing, while the side chains of Cys966, Asn998, Asn1008, and Gln1029 are shown as ball-and-stick drawings. Thick dotted lines represent hydrogen bonds and numbers show the O…N distances in angstroms. Thin dotted lines show the positions where the structural changes are expected to occur.


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Supporting Information The following files are available free of charge. Difference FTIR spectra for native and uniformly 15N-labeled neo1-LOV2 in the 3000–2730 cm– 1

region (Figure S1, PDF). Difference FTIR spectra for Arabidopsis phot1-LOV2 and phot2-

LOV2, and Adiantum neo1-LOV2 and neo1-LOV1 in the 3000–2760 cm–1 region (Figure S2, PDF).

Corresponding Author * To whom correspondence should be addressed. Phone & FAX: 81-52-735-5207. E-mail: [email protected]

Present Addresses #

T. I.: Department of Pharmaceutical Science, Toho University, Funabashi, Chiba 274–8510,

Japan

Funding Sources This study was supported by grants from the Japanese Ministry of Education, Culture, Sports, Science, and Technology to H. K. (25104009, 15H02391), M. U. (26410017), and T. I. (16K07318).

Some of the computations were performed at the Research Center for

Computational Science, Okazaki, Japan.


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Acknowledgements The authors would like to thank Enago (www.enago.jp) for the English language review.

ABBREVIATIONS LOV, light-oxygen-voltage; PAS, Per-Arnt-Sim; FMN, flavin mononucleotide; FTIR, Fourier transform infrared.


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[36] Kottke, T., Heberle, J., Hehn, D., Dick, B., and Hegemann, P. (2003) Phot-LOV1: photocycle of a blue-light receptor domain from the green alga Chlamydomonas reinhardtii, Biophys J. 84, 1192–1201. [37] Zhu, J., Mathes, T., Hontani, Y., Alexandre, M. T., Toh, K. C., Hegemann, P., and Kennis, J. T. (2016) Photoadduct Formation from the FMN Singlet Excited State in the LOV2 Domain of Chlamydomonas reinhardtii Phototropin, J. Phys. Chem. Lett. 7, 4380–4384. [38] Kasahara, M., Swartz, T. E., Olney, M. A., Onodera, A., Mochizuki, N., Fukuzawa, H., Asamizu, E., Tabata, S., Kanegae, H., Takano, M., Christie, J. M., Nagatani, A., and Briggs, W. R. (2002) Photochemical properties of the flavin mononucleotide-binding domains of the phototropins from Arabidopsis, rice, and Chlamydomonas reinhardtii, Plant Physiol. 129, 762– 773. [39] Iwata, T., Nozaki, D., Tokutomi, S., and Kandori, H. (2005) Comparative investigation of the LOV1 and LOV2 domains in Adiantum phytochrome3, Biochemistry 44, 7427–7434. [40] Iwata, T., Tokutomi, S., and Kandori, H. (2002) Photoreaction of the cysteine S-H group in the LOV2 domain of adiantum phytochrome3, J. Am. Chem. Soc. 124, 11840–11841. [41] Ataka, K., Hegemann, P., and Heberle, J. (2003) Vibrational spectroscopy of an algal Phot-LOV1 domain probes the molecular changes associated with blue-light reception, Biophys J. 84, 466–474. [42] Bednarz, T., Losi, A., Gartner, W., Hegemann, P., and Heberle, J. (2004) Functional variations among LOV domains as revealed by FT-IR difference spectroscopy, Photochem. Photobiol. Sci. 3, 575–579.


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For Table of Contents Use Only

Hydrogen Bonding Environment of the N3–H Group of FMN in the LOV Domains of Phototropins

Tatsuya Iwata,†,# Dai Nozaki,† Atsushi Yamamoto,† Takayuki Koyama,† Yasuzo Nishina,‡ Kiyoshi Shiga,§ Satoru Tokutomi,|| Masashi Unno,⊥ and Hideki Kandori*,†


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Hydrogen Bonding Environment of the N3âH Group of Flavin

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