Vibrational Mode Analysis of Isotope-Labeled Electronically Excited

ARTICLE pubs.acs.org/JPCB

Vibrational Mode Analysis of Isotope-Labeled Electronically Excited Riboflavin Matthias M. N. Wolf,† Herbert Zimmermann,‡ Rolf Diller,*,† and Tatiana Domratcheva*,‡ † ‡

Fachbereich Physik, TU Kaiserslautern, D-67663 Kaiserslautern, Germany Max-Planck-Institut f€ur medizinische Forschung, D-69120 Heidelberg, Germany

bS Supporting Information ABSTRACT: Isotope-labeled riboflavin in DMSO was employed in conjunction with femtosecond time-resolved infrared vibrational spectroscopy and quantum chemical calculations to analyze and assign the electronically excited state vibrational modes of the isoalloxazine unit as a prototype for the cofactors in flavin binding blue-light receptors. Using the riboflavin 13Canalogues RF-2-13C and RF-4,10a-13C, the carbonyl vibrations, in particular, were studied. Various quantum chemical models were applied that take into account a polarizable environment or the impact of hydrogen bonds. The CIS quantum-chemistry method was successfully applied to describe the lowest singlet excited electronic state in riboflavin. The experimentally observed frequencies and isotope-shifts as well as their variability in the diverse model calculations are discussed. On these grounds, a consistent assignment of the electronic ground and excited state vibrations is presented.

’ INTRODUCTION Flavins serve as photoactive prosthetic groups in blue-light receptors mediating diverse light responses such as phototropism, chloroplast relocation, stomata opening, growth inhibition, photoperiodism, and phototaxis in plants, bacteria, green alga and fungi.1 Families of flavin-containing blue-light receptors identified so far include light-oxygen-voltage (LOV) domaincontaining proteins, cryptochromes and blue light using flavin adenine dinucleotide (FAD) proteins (BLUF). There is no unique flavin photoreaction activating the blue-light receptors, therefore, each class is based on its own photochemistry as determined by the interaction of the flavin with the specific residues in its binding pocket. Flavins are capable of one- and two-electron reduction, e.g., from Tyr, Trp, or Cys residues, and their ability to accept an electron is enhanced upon electronic excitation. Irradiation of the flavin chromophore, e.g., riboflavin (RF), flavin mononucleotide (FMN) or flavin adenine dinucleotide (FAD) around 450 nm results in electronically excited singlet or triplet states of the isoalloxazine ring system (Scheme 1). The diverse subsequent primary reactions that finally lead to the respective signaling state embrace formation of a cysteinyl-flavin adduct in LOV,2
4 one-electron reduction of the oxidized flavin in cryptochromes5,6 or the rearrangement of a hydrogen-bonded network among FAD and nearby amino acid side chains in BLUF.7
13 Identification and characterization of the various coupling mechanisms and their dynamics is an important part of current blue-light receptor research. Ultrafast infrared spectroscopy has widely been used to incisively probe the electronically excited cofactor and its interactions with the protein on structural grounds, e.g., in retinal proteins14,15 and phytochrome.16 Recently, IR spectra of the r 2011 American Chemical Society

singlet excited flavin chromophore were reported for the LOV17 and BLUF11,13 photoreceptors, FAD in D2O18 and RF in DMSO.19 Vibrational modes with local O
H, N
H, CdO or CdN stretching character are well suited to probe the dynamics of hydrogen bonding and thus the structural dynamics during the transient states of the primary photoreaction, especially in conjunction with the protein crystal structure. Prerequisite for this is a detailed vibrational mode analysis of the electronic states involved. In the electronic ground state, a classical and wellproven approach is to experimentally characterize the vibrational spectra of the cofactor (here the isoalloxazine group as, e.g., in RF, including isotopologues) on the one hand, and on the other hand to apply quantum chemical normal-mode analysis. In the electronically excited state, this approach represents a challenge from both experimental and computational point of view. Following this strategy we already studied photoinduced vibrational dynamics of RF in DMSO in conjunction with the configuration interaction singles (CIS) excited electronic state calculations.19 Spectral changes occurring on a 4 ps time scale were assigned to vibrational cooling/relaxation of the isoalloxazine chromophore in the excited singlet ππ* state enabling identification of excited state vibrational bands. Quantum chemical calculations of riboflavin in the first excited singlet state indicated a loss of the double bond between N5 and C4a. Upon ππ* electronic excitation, pronounced frequency down-shifts were found for double-bond stretching modes, e.g., CdO, CdC, and CdN of the isoalloxazine ring. Received: November 11, 2010 Revised: April 7, 2011 Published: May 20, 2011 7621

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Scheme 1. Structure of Riboflavin

In this paper we study for the first time the isotope induced shifts of vibrational modes in electronically excited riboflavin, in particular that of the carbonyl vibrations that potentially report on intra- and intermolecular interactions, e.g., hydrogen bonding, solvent polarization as well as the flavin redox potential and redox state.20 We present electronic ground and first excited state vibrational spectra of two isotope labeled compounds, 2-13Criboflavin (RF-2-13C) and 4,10a-13C-riboflavin (RF-4,10a-13C). Our previously published normal-mode analysis is extended to include isotopic shifts. In addition, we present several new models to account for carbonyl-solution interaction.

’ MATERIALS AND METHODS Transient IR Spectroscopy. The UV-pump/mIR-probe experiments were performed as described earlier:19 The output of a Ti:Sa kHz amplifier (CPA 2001, Clark-MXR) at 775 nm with 150 fs duration (full width at half-maximum, fwhm) was used to generate pump pulses at 387 nm by second harmonic generation and probe pulses by two-stage optical-parametric-amplification with consecutive difference frequency generation (350 fs fwhm of pump/probe cross correlation). Synthesis of Isotopically Labeled Riboflavins RF-2-13C and RF-4,10a-13C. 1-(D-ribitylamino)-3,4-dimethylbenzene was coupled with phenyldiazonium chloride to obtain 1-(ribitylamino)-2-phenylazo-4,5-dimethylbenzene.21 By reacting the azo-compound with 2-13C- or 4,6-13C2-barbituric acids, the labeled riboflavins were prepared. The labeled acids were synthesized by condensing 13Curea with diethylmalonate using sodiumethoxide as the condensing agent or by reacting urea with 1,3-13C2 diethylmalonate. The labeled riboflavins migrated as one spot in silica (TLC, 5CHCl3:5CH3OH:2 ethyl acetate). The precise labeling (99% 13C) and chemical purity (by 99%) were confirmed by the 13C NMR and mass spectra. Both compounds were dissolved in DMSO-d6 (99.8%, Merck) at a concentration of 13 mM, as in our earlier experiments.19 The sample was placed between two CaF2 windows with a 200 μm PTFE spacer. During measurement the sample was rotated to exchange the excited sample volume between consecutive laser shots. To avoid respectively control photodegradation steady state UV/vis and FTIR transmission spectra were recorded regularly. FTIR spectra of employed samples are shown in Figure 1. Quantum-Chemical Calculations. Optimized geometries, vibrational frequencies and isotopic shifts for a number of flavin models were computed by means of the B3LYP/6-31G(d) and CIS/6-31G(d) methods in the ground and excited electronic states, respectively. Calculations were performed by using the Gaussian03 quantum-chemistry package.22 The polarizable continuum model (PCM)23 with the default DMSO parameters was employed to model the bulk solvent effects.

Figure 1. FTIR spectra of investigated riboflavin compounds with solvent (DMSO-d6) spectra subtracted. Note that the isotope label induced band-shifts, especially in the carbonyl region.

’ RESULTS AND DISCUSSION Infrared difference absorption spectra of RF-4,10a-13C and RF-2-13C were obtained after excitation at 387 nm between 1720 and 1300 cm
1 for delay times up to 200 ps (Figure 2). The difference spectra show large positive and negative bands, corresponding to excited state and ground state vibrational bands, respectively. Significant spectral shifts are evident when comparing both isotopically labeled compounds with the unlabeled riboflavin, especially in the carbonyl region. On a time scale of several picoseconds small absorption changes are visible, associated with a slight blue-shift of positive absorption maxima. The data were fitted to a multiexponential model as described before:19 ΔODðλ, tÞ ¼ A0 ðλÞ þ

n

Ai ðλÞ 3 e
1=τ ∑ i¼1

with time constants τi and amplitudes Ai(λ) (decay associated spectra, DAS), where A0(λ) represents the difference spectrum after long delay times (see Supporting Information). The DAS of the isotope labeled compounds in comparison with our previously published data of unlabeled riboflavin are shown in Figure 3. For both isotopologues two exponentials were sufficient to fit the data. The first time constant is similar to the previously reported 4.8 ps component in unlabeled RF: τ1 = 4.6 ( 0.2 ps for RF-2-13C and 4.9 ( 0.2 ps for RF-4,10a-13C. Again,19 by its characteristic spectral features this process is assigned to vibrational cooling/relaxation in the S1 state. Accordingly, pairs of negative and positive lobes correspond to the vibrationally relaxed and unrelaxed state, respectively (cf. Figure 3d). The second time constant τ2 = 122 ps (RF-2-13C) and 124 ps (RF-4,10a-13C) has not been found in the dynamics of unlabled riboflavin in DMSO. In Figure 3, the spectral features of this component (A2) does not show general spectral correlation with either A1 or A0. The physical nature and significance of A2 is unclear at this time and will be subject of future studies, however, it does not interfere with the vibrational analysis presented in this work. Component A0 essentially shows the difference between the S0 and S1 state of the respective riboflavin derivate. Negative and positive bands correspond to S0 and S1 vibrations, respectively. Band positions were determined with a Lorentzian lineshape fit (see markers in Figure 3, Table 1 and Supporting Information, Tables S1
S4). In addition, the relaxed S1 vibrational 7622

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Figure 2. Difference spectra of (a) RF-4,10a-13C and (b) RF-2-13C in comparison with (c) unlabeled riboflavin19 at selected delay times.

Figure 3. Decay-associated-spectra (DAS) of (a) RF-4,10a-13C and (b) RF-2-13C in comparison with (c) unlabeled Riboflavin.19 Enlarged A1 are shown in part d. Red markers show the spectral position of difference bands as found with a Lorentzian line-shape fit (description see Supporting Information).

frequencies can be identified by the respective negative lobe of the characteristic pairwise pattern in A1, especially when S1 bands are obscured by strong and overlapping S0 features. To aid the vibrational assignments of the isotope labeled riboflavin, normal-mode analysis for the electronic ground state and the first excited singlet state, which has a ππ* character,19 was performed on the grounds of various lumiflavin and riboflavin models presented in Figure 4. Lumiflavin (LF) represents the simplest model of the flavin chromophore. Riboflavin models RF-HB and RF were used in our previous work for theoretical assignments of riboflavin vibrations.19 Model RF-HB accounts for a stabilizing intramolecular hydrogen bond in the riboflavin molecule. Model lumiflavin
water (LF-W) contains a similar hydrogen bond and predicts the vibrations of the isoalloxazine ring which are very close to those of the RF-HB model. Riboflavin
water models (RF-HB-W-1 and RF-HB-W-2) account for hydrogen bonding interactions of C2dO2 with a hydroxyl group and of C4dO4, N3
H or N5 with a water molecule

(see Scheme 1 for riboflavin atom numbering). Models LF, LF-W, RF, RF-HB, RF-HB-W-1, and RF-HB-W-2 depicted in Figure 4 are referred in the following as molecular models. In addition to the molecular models, two DMSO solvent models, LF-DMSO and LF-W-DMSO, were used to address the bulk solvent effect by embedding the LF and LF-W molecules in polarizable continuum23 representing the DMSO solvent. The normal mode vibrational analysis was carried out for the three isotopologues of each model. The computed frequencies, IR intensities and the isotopic shifts in the spectral region of interest are collected in the Supporting Information, Tables S1
S4. As demonstrated in our previous work,19 the vibrations characterized by noticeable IR intensities typically belong to the isoalloxazine chromophore and not to the ribityl chain of the riboflavin molecule. This finding was extensively used in the theoretical assignments of the riboflavin photodynamics.19 In this work, in addition to the good correspondence of the computed normal-mode frequencies and intensities with the 7623

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Table 1. Experimentally Determined Vibrational Frequencies with Corresponding Isotopic Shifts (cm
1) Indicated in Brackets (2-13C and 4,10a-13C) Compared to the Computational Results of the Two Representative Quantum-Mechanical Models LFDMSO and RF-HB-W-2a LF-DMSO exptl freq, cm


1

calcd freq, cm


1

RF-HB-W-2

normal mode as atomic displacements

calcd freq, cm


1

normal mode as atomic displacements

Ground State 1711 (
1;
40)

1771 (
3;
26)

C4, O4, H3, C2, O2

1807 (þ1;
43)

C4, O4, H3

1676 (
47;
5)

1734 (
37;
19)

C2, O2, H3, C4, O4

1771 (
45;
1)

C2, O2, H3

1635 (
1;
11)

1584 (0;
4)

1618 (
0;
4)

N5, C4a, C10a, N1

1547 (0;
7)

1582 (
1;
5)

N10, C10a, C4a, C9,

1511 (0;
18)

1560 (
1;
20)

1464 (þ1;
5)

1526 (0; 0)

1591 (0;
9)

N1, N5 N5, C10a, N1, H(CH3)

C8, C9, C9a, C5a, N10, N5, C4a, C10a, N1 C5a, C6, C7, C8, C9, C9a, C10, C10a, N1

1575 (0;
16)

C8, C9, C9a, N10, N5, C4a, C10a, N1

1541 (0;
3)

H(CH3), H9, H6, C5a, C6, C7, C8, C9, C9a, C10a, C4a

1516 (0;
1) 1427 (
4;
1) 1345 (
5;
14)

1302 (--;
5)

1475 (
1;
4)

C7, C8, C9, C6, C5a, C10a,

1396 (
3;
12)

N1, C2, N3, C4 C8, C9, C7, C6, C4, N3, C2

1368 (
5;
13)

1521 (0; 0) 1478 (
1;
3)

C8, C9, C9a, C5a, C4a

1425 (
3;
8)

1332 (0;
4) 1278 (--; --)

H(CH3), C4a, N5, C9a, C5a

H(CH3), H9, C9a, C5a, C10a, C4a H(CH3), C5a, C7, C8, C9a, N10, C4a

1439 (
1;
2)

H(CH3), H9, H6, C10a, C4a, N1

1382 (
3;
6)

H(rib), H6, N10, C10a, C2, H3

H6, C7, C8,C9a,N10, C10a, C2, N3, C4

1367 (
4;
6)

H6, H(rib), C5a, C9, C10a, C2, C4

C6, C5a, C9a, N10, N5, N1, N3, C4

1342 (
1;
4)

C5a, C9a. N10, N5, C4, N3,C2

1328 (
1;
2)

N5, N10, H9, H6, C5a, C9a

1303 (
2;
1)

H6, H9, C5a, N5

1334 (0; 0) 1304 (
1;-2) 1289 (0; 0)

H6, H9, C9a, C5a, N5, N10, C4a H6, H9, C5a, C7, N5, N10, C2, H3 H(rib), C9a, N10, C5a, N5, C4a, C2, N3. C4

Excited State 1652 (
1;
44) 1642 (
44;
5)

1906 (
8;
62) 1877 (
37; þ13)

1571 (
1;
3)

1767 (
1;
14)

1546 (
4;
44)

1748 (
1;
8)

1501 (
1; þ1)

1709 (0;
9)

1963 (
2;
49) 1953 (
47; þ1)

C4, O4 C2, O2, H3

C4, O4, H3, C2, O2 C2, O2, H3, C4, O4

C6, C7, C5a, N5,C4a,C10a,N1

1771 (0;
12)

C6, C7, C8, C9, C4a, C10a, N1

C6, C7, C8, C9, C9a, C5a,C10a, N1

1750 (0;
25)

C6, C7, C8, C9, C9a, C4a, C10a, N1

1687 (0;
1)

C6, C7, C9a, C5a, H6, H9, H(CH3)

1659 (0;
1)

C5a, N5, C4a, C6, H6, H9, H(CH3)

C5a, C6, C7, C8, C9, C9a, N10, N5, C10a, N1

1454 (
7; 0)

1674 (0;
4)

C5a, C7, C8, C9a, N5, N10, C4a, C10, N1

1648 (0,
1)

H6, H9, H(CH3), C6, C7, C8, C9,

1641 (0, 0) 1413 (
8;
12)

C9a, C5a, C10a, C4a H(CH3), C7, C9, C5a, N10

1600 (
1;
4)

H6, H9, H(CH3),C6, C9,

1582 (
4;
5)

H(CH3), H3, C4, C4a, N3, C2, C5a,

1618 (0; 0)

H(CH3), C4a, N5

1607 (
1;
3)

H9, H6, H(CH3), C9a, N10, N5a,

1593 (
2;
3)

H(rib), C4a, C10a, N10

C9a, N10, C10a,N5, C4a, N1, H3

C10a, C4, N1, C4a

C9a, C7, C8 1567 (
0;
1) 1376 (
3;
12)

1532 (
6;
6)

H3, H9, C7, C8, C9, C9a, C5a, C10a,

1540 (0; 0)

H(CH3), H(rib), H3, N3 H3, C2, N3, N1, C10a, C4a, C5a, C6, C7, H9

C4a, C4, N3, C2, C1 1539 (
4;
4) 1290 (--; --)

1458 (
1;
5)

1273 (--; --)

1437 (0;
3)

C7, C8, C9, C9a, C5a, N10, C10a, C4a, N5 C5a, C6, C7, C8, C9, C9a, N10, N5,

H3, C4a, C10a, C2, N3, N10, C9a, C5a

1527 (
5;
8)

H(rib), C10a, N10, C4a, N5, C4, N1, N3, C2

1467 (
2;
6)

H(rib), N10, C10a

1436 (0; 0)

H6, H9, C5a, C6, C7, C9, C9a, N10, N5, C4a

C10a, C4a, C4 1412 (
1;
4)

H6, H9, C9a, C5a, C10a, C4a, N3, C2

1415 (
1;
6)

1225 (--; --)

1370 (0;
2)

C5a, C6, C7, C8, C9, C9a, N10, C10a

1370 (0; 0)

1182 (--; --)

1345 (
10;
11) 1324 (
6;
2)

C4a, C2, N3, C4, N1, C6, C7, C8, C9 H6, C7, C7, C9, C9a, C4a, C4, C2, N3

1346 (
2;
3) 1342 (
12;
10) 1318 (0; 0)

H(rib), H9, C6, C5a, C9a, C8, N10, C10a, C4a H6, H9, C5a, C6, C7, C9, C9a, H(rib) H6, C5a, C6, C7, C9a, C10a, C4a, C2 H6, H9, C7, C8, C4a, C4, N3, C2, N1, H3 H(rib), C7, C8, C6, C9, N10, N3, C2, C4

a

The calculated frequencies are given without scaling. A complete set of computed frequencies in the spectral region of interest is given in the Supporting Information, Tables S1
S4. 7624

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Figure 4. Molecular models used in the quantum-chemical normalmode analysis (cf. text).

experimentally observed bands, we rely on the computed and experimental isotopic shifts. For model LF DMSO, the computed IR spectra of the three isotopologues are presented as stick spectra in Figure S1 in the Supporting Information. The assignment of the experimentally determined vibrational frequencies to the computed normal modes is presented in Table 1 where two representative models, i.e., LF-DMSO and RF-HB-W-2, are used. The correlation between the experimental and computed (without scaling) frequencies in Table 1 is shown in Figure 5. All molecular models overestimate the carbonyl CdO stretching frequencies. Figure 5 clearly demonstrates this disagreement for model RF-HB-W-2. In our previous work, to explain the disagreement we proposed that additional hydrogen bonds, which were not considered in model RB-HB, were formed in the studied samples.19 However, in this study, model RF-HB-W2 yielded the up-shifted frequencies of the carbonyl stretches as well. Interestingly, including bulk polarization using a polarizable continuum model in LF-DMSO led to a downshift of the carbonyl frequencies thus improving agreement between calculations and experiment. Hence, the downshift of the carbonyl stretching frequencies noted in DMSO in our previous study19 should be assigned to the bulk solvent effect (solvation) rather than to the specific hydrogen-bonding interactions. In line with this assignment, the effect of an additional hydrogen bond on the carbonyl downshift in model LF-W-DMSO is insignificant (Supporting Information, Tables S1 and S3). Ground-State Vibrations. The 1711 and 1676 cm
1 bleach bands in the light-induced difference spectra were assigned to the C4dO4 and C2dO2 carbonyl stretching vibrations, respectively, of the ground state riboflavin.19 The assignments of these bands (further referred as the higher- and lower-frequency carbonyl bands, respectively) specifically to the C4dO4 and C2dO2 stretches were complicated by the discrepancies between the experimental and computed frequencies in our previous work. Isotopic labeling and new quantum-chemical models allow us now to unambiguously assign the carbonyl stretches. In the RF-4,10a-13C spectrum (Figure 3a), we observe that the bleach around 1711 cm
1 disappears, while the other bleach, now at 1671 cm
1, grows stronger and broader, in line with the FTIR spectra (Figure 1). A shift of any carbonyl mode to an even lower frequency would coincide with the strong (positive) product feature at 1637 cm
1 (Figure 3a) and reduce its apparent

Figure 5. Correlation between the experimental and normal-mode frequencies in Table 1 demonstrating the quality of the assignments: top, electronic ground state; bottom, electronic excited state; red circles, model RF-HB-W-2; black squares, model LF-DMSO; solid line, linear fits for LF-DMSO. The overestimated carbonyl stretching frequencies (cf. text) are indicated.

intensity, which is not observed. It is thus concluded that the mode at 1711 cm
1 downshifts to approximately 1671 cm
1, thereby strongly overlapping with the 1676 cm
1 mode that shows no significant down-shift (approximately 5 cm
1). Accordingly, we assign the 1711 cm
1 frequency to the C4dO4 stretch. In the RF-2-13C spectrum (Figure 3b), the 1711 cm
1 band does not shift, while the bleach at 1676 cm
1 disappears. A new band appears at 1629 cm
1 which can be assigned to the labeled 13C2dO2 stretch and is also observed in the FTIR spectra of the ground state (cf. Figure 1). Alternatively, 1629 cm
1 can correspond to the highest-frequency CdC stretching mode, which is usually observed at 1620
1630 cm
1 and is obscured by the strong S1 positive absorption in unlabeled riboflavin19 (Figure 3c). However, the highest CdC mode is low in intensity (see Figure 1 and ref 24). Therefore, the 1629 cm
1 band is assigned to the labeled 13C2dO2 stretch. The assignment of the 1711 and 1676 cm
1 bands to the C4dO4 and C2dO2 stretch, respectively, is in agreement with the results of previous labeling experiments.20,25 Our quantumchemical calculations clearly indicate that only one CdO stretching frequency is affected by the 13C2- or 13C4-labeling. Moreover, the isotopic shift is about the same magnitude as the 7625

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The Journal of Physical Chemistry B difference between the two carbonyl peaks. Therefore, the downshifted upper frequency coincides with the lower frequency, as it is observed in the 4,10a-13C-labeled spectrum. Upon 4,10a-13C-labeling, the experimentally determined shift of the higher-frequency mode of approximately 40 cm
1 is in good agreement with the computed shifts of 40
45 cm
1 in the molecular models. Upon 2-13C-labeling, our models predict a downshift of 36
46 cm
1 for the lower carbonyl frequency. The experimentally determined downshift of 47 cm
1 could be overstated due to the close-by highest ring CdC stretching mode and the product (excited state) bands. Previously, a smaller approximately 30 cm
1 2-13C shift has been reported for the lower-frequency carbonyl stretch of solid lumiflavin.24,26 It is interesting to mention that the DMSO models predict smaller carbonyl isotopic shifts. In addition, the lower carbonyl frequency is predicted to show 2-13C as well as 4,10a-13C downshifts, therefore suggesting considerable coupling of the two carbonyl vibrations. Notably, the coupling of the carbonyl stretches is not observed in the experimental spectra: Upon 2-13C and 4,10a-13C labeling only one of the carbonyl stretching bands, respectively, is found downshifted. As noted above, the polarization by DMSO was important to predict the positions of the carbonyl stretching bands. In contrast, the isotopic shifts are predicted fairly well by the molecular models. Thus, the observed ground-state isotopic shifts can be interpreted without involving the bulk solvent effect (solvation). The RF-4,10a-13C spectrum can serve to study the electronic structure of the flavin CdC/CdN double bonds as several corresponding bands were found downshifted. The 1584 cm
1 band downshifts by 4 cm
1, the 1547 cm
1 band by 7 cm
1, whereas the 1511 cm
1 by 18 cm
1. These three bands are assigned to the stretches of the N5dC4a-C10adN1 π-conjugated electronic system. The respective computed shifts are in reasonable agreement with the experiment (see Supporting Information, Tables S1
S2). In previously reported FTIR20 and Raman25 spectra of the isotope-labeled FADs in water and D2O, similar features for the double-bond CdC and CdN stretching frequencies were found. We do not observe any effect of the 13C2-labeling in the CdC/CdN double-bond spectral region. Instead there are small downshifts of the lower frequencies at 1427 and 1345 cm
1, assigned to single-bond stretching modes which are rather delocalized over the flavin ring system. In addition, single-bond stretches at 1464, 1345, and 1302 cm
1 are affected by the 4,10a-13C-labeling. Excited State Vibrations. The positive features of the A0 spectrum were assigned to the first ππ* excited singlet state of riboflavin:19 The derived frequencies are in agreement with the previously reported S1 vibrations of the flavin chromophores in the LOV and BLUF photoreceptors.13,17 Light excitation invokes characteristic changes in the CdN and CdC double-bond stretching bands. In addition, pronounced downshifts of the carbonyl stretches to 1652 and 1642 cm
1 were observed in RF in DMSO.19 Note that the difference between the excited carbonyl stretch frequencies is only 10 cm
1 which is smaller than the expected 13C isotopic shifts of about 40 cm
1, suggesting that the downshifted higher carbonyl frequency could become the lower frequency in the isotope-labeled spectrum. In contrast to the electronic ground state, there is a disagreement among molecular models about the carbonyl frequency isotopic shifts in the excited state and, therefore, about the assignments of these frequencies to the C2dO2 and C4dO4 stretches, indicating high sensitivity of these band-positions to

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the specific environment and interactions of the uracile ring III. The CdO isotopic shifts predicted by the LF, RF and RF-HB-W1 models (see Supporting Information, Tables S3 and S4) are at odds with the other models and experiment. Therefore, we did not use these models for the assignments of the excited state vibrations. Best agreement between the computed and observed shifts was obtained with the molecular model RF-HB-W-2 (see Table 1). Both DMSO models predict the right trend but somewhat overestimate the isotopic shift of the C4dO4 as compared to that of the C2dO2 stretch. In addition, these models predict a significant upshift of the lower frequency carbonyl band upon the 13C4,13C10a-labeling which is in disagreement with the experiment. In the RF-2-13C spectrum (Figure 3b), the higher-frequency carbonyl stretch at 1652 cm
1 does not shift while the lowerfrequency band shifts to 1598 cm
1, suggesting that the latter corresponds to the C2dO2 vibration. In the RF-4,10a-13C spectrum (Figure 3a), two new bands appear at 1637 and 1611 cm
1. This could imply two possibilities: The original lower-frequency band at 1642 cm
1 is slightly downshifted, while the higherfrequency 1652 cm
1 band is downshifted by about 40 cm
1. Alternatively, both bands could be downshifted by 15 and 31 cm
1 respectively. Taking into account the 2-13C isotopic shift, we favor the first possibility which is also supported by the calculated shifts. Thus, the 1652 and 1642 cm
1 bands are assigned to the C4dO4 and C2dO2 stretching modes of the exited S1 flavin, respectively. For this assignment, the magnitudes of the experimental isotopic shifts for the (negative) ground state and (positive) excited state CdO bands are in good correspondence (see Table 1). The ππ* excitation reduces the double-bond character of the isoalloxazine ring, which can be recognized in the characteristic changes of the double-bond stretching region. In the ground state, the double-bond stretches are assigned to bands at 1620
1630 cm
1 (CdC ring I, obscured by the positive absorption in Figure 3c), 1584, 1547, 1511, and 1464 cm
1 (mixed CdC and CdN). In the excited state, the double-bond stretching vibrations are at 1571 cm
1 (CdC, C10adN1), 1546 cm
1 (C10adN1), 1501 and 1454 cm
1 (mixed CdC and CdN). These bands can be regarded as marker bands of the isoalloxazine excited electronic structure; their assignments in Table 1 are supported by the agreement between computed and experimentally determined isotopic shifts. In Supporting Information, Figure S2 demonstrates the structure of the first excited ππ* state and the normal modes assigned to the marker bands. As the 1547 (S0) and 1546 cm
1 (found in S1) features obviously coincide (Figure 3c), the latter could not be observed in the difference spectra of the photoexcited LOV and BLUF photoreceptors.13,17 We could identify the 1546 cm
1 feature of the excited state in unlabeled riboflavin in DMSO by an asymmetry in the strong 1547 cm
1 bleach, supported by the S1 cooling feature of A1 at this spectral position.19 In the RF4,10a-13C spectrum (Figure 3a) the cooling feature in A1 is missing, and the asymmetry is not found. The possibility for the downshifted 1546 cm
1 band is the cooling feature in A1 at 1502 cm
1, which coincides with the 1501 cm
1 band of the unlabeled compound. Therefore, the band at 1546 cm
1 either disappears or downshifts by more than 40 cm
1 to coincide with the 1502 cm
1 peak in RF-4,10a-13C, which, according to the calculations, undergoes only minor shift of approximately 1 cm
1. The large isotopic shift suggests that the 1546 cm
1 corresponds to a rather localized vibration. 7626

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The Journal of Physical Chemistry B On the basis of the normal-mode analysis, 1546 cm
1 can be assigned to a localized C10adN1 double-bond stretching. Our models predict a downshift of this frequency by 25
29 cm
1 in the 13C4,13C10a-labeled flavin. Note that none of the models yields a shift as large as 40 cm
1. The DMSO models predict a modest downshift of 8 cm
1 together with the downshift of the 1571 cm
1 mode by 14 or 16 cm
1, corresponding to a rather delocalized vibration. In the excited state the N5dC4a double bond is significantly weakened and its stretching frequency is significantly lowered, therefore it does not contribute to the 1546 cm
1 vibration. In RF-2-13C the 1546 cm
1 band downshifts 4 cm
1 (experimental) with practically no shift according to our calculations. Upon 13C2-labeling, the 1454 and 1413 cm
1 bands downshift by 7 and 8 cm
1, respectively, whereas the bands at 1413 and 1376 cm
1 downshift by 12 cm
1 in the RF-4,10a-13C spectrum. The observed changes are in line with the previous assignments of these bands to the stretching vibrations of the uracile ring III.19 Our current calculations predict somewhat smaller shifts than the observed ones. However, an overall agreement of calculated and experimental shifts could be noticed. The 1413 and 1376 cm
1 frequencies are assigned to the delocalized modes with the N10
C10a and C4a-C10a contributions, respectively. The 1376 cm
1 band corresponds to the 1345 cm
1 band in the ground state; its upshift in the excited state may be related to the strengthening of the C4
C10a bond upon electronic excitation as the structure of the first excited ππ* state19 suggests (see Figure S2 in Supporting Information). In conclusion, ultrafast transient infrared spectroscopy combined with quantum chemical calculations is a powerful tool to identify and analyze vibrational modes, i.e., molecular structure of the transient electronic states of photoexcitable molecules as, e.g., flavins and analogues. Identification of the involved excited electronic states and their spectroscopic markers is critical for understanding the photodynamics and chemical reactivity in biologically relevant chromoproteins. In a first step along this line, we earlier presented the results on excited state vibrational dynamics of nonlabeled riboflavin in DMSO together with normal-mode analysis.19 Here, we extended this work to the isotope-labeled riboflavins. Labeling allowed assignment of the CdO carbonyl stretching frequencies as marker bands for interactions and the CdC/CdN double bond stretches as markers for the electronic structural change. The CIS method provides satisfactory explanation of the excited state vibrational spectra and the isotopic shifts. Including the DMSO polarization effect by means of the PCM model improves the predicted frequencies of the carbonyl vibrations. The DMSO models on the other hand tend to predict a higher delocalization of vibrational modes than the experimental shifts suggest. In addition, a new picosecond kinetic component was found in both isotope-labeled flavins. This component could be a manifestation of the mass-effect in the excited state dynamics, which cannot be assigned to a specific process yet and requires further studies.

’ ASSOCIATED CONTENT

bS

Supporting Information. Description of data evaluation, Figure S1 showing the computed IR spectra of the three isotopologues of LF-DMSO in the electronic ground and excited states, Figure S2 presenting the normal modes assigned to the marker bands and the electronic structure of the ground and first excited singlet ππ* state, and Tables S1
S4 containing the computed vibrational frequencies, IR intensities and isotopic shifts of various

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flavin models in the ground and excited states compared to the experimental vibrational frequencies. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*(R.D.) E-mail: [email protected]. Telephone: 49-631205-2323. Fax: 49-631-205-3902. (T.D.)E-mail: Tatjana. [email protected]. Telephone: 49-6221486-504. Fax: 49-6221-486-585.

’ ACKNOWLEDGMENT H.Z. and T.D. are very grateful to Ilme Schlichting for constant support. M.M.N.W. and R.D. acknowledge support by OTLAP Forschungsschwerpunkt Rheinland-Pfalz and Stiftung Rheinland-Pfalz f€ur Innovation. ’ REFERENCES (1) Briggs, W. R.; Spudich, J. L. Handbook of photosensory receptors; Wiley-VCH: Weinheim, Germany, 2005. (2) Christie, J. M. Annu. Rev. Plant Biol. 2007, 58, 21. (3) Swartz, T. E.; Corchnoy, S. B.; Christie, J. M.; Lewis, J. W.; Szundi, I.; Briggs, W. R.; Bogomolni, R. A. J. Biol. Chem. 2001, 276, 36493. (4) Kottke, T.; Heberle, J.; Hehn, D.; Dick, B.; Hegemann, P. Biophys. J. 2003, 84, 1192. (5) Bouly, J. P.; Schleicher, E.; Dionisio-Sese, M.; Vandenbussche, F.; Van Der Straeten, D.; Bakrim, N.; Meier, S.; Batschauer, A.; Galland, P.; Bittl, R.; Ahmad, M. J. Biol. Chem. 2007, 282, 9383. (6) Berndt, A.; Kottke, T.; Breitkreuz, H.; Dvorsky, R.; Hennig, S.; Alexander, M.; Wolf, E. J. Biol. Chem. 2007, 282, 13011. (7) Masuda, S.; Hasegawa, K.; Ono, T. Biochemistry 2005, 44, 1215. (8) Gauden, M.; van Stokkum, I. H. M.; Key, J. M.; L€uhrs, D. C.; Van Grondelle, R.; Hegemann, P.; Kennis, J. T. M. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 10895. (9) Jung, A.; Reinstein, J.; Domratcheva, T.; Shoeman, R. L.; Schlichting, I. J. Mol. Biol. 2006, 362, 717. (10) Gauden, M.; Yeremenko, S.; Laan, W.; van Stokkum, I. H. M.; Ihalainen, J. A.; van Grondelle, R.; Hellingwerf, K. J.; Kennis, J. T. M. Biochemistry 2005, 44, 3653. (11) Stelling, A. L.; Ronayne, K. L.; Nappa, J.; Tonge, P. J.; Meech, S. R. J. Am. Chem. Soc. 2007, 129, 15556. (12) Domratcheva, T.; Grigorenko, B. L.; Schlichting, I.; Nemukhin, A. V. Biophys. J. 2008, 94, 3872. (13) Bonetti, C.; Mathes, T.; van Stokkum, I. H. M.; Mullen, K. M.; Groot, M.-L.; van Grondelle, R.; Hegemann, P.; Kennis, J. T. M. Biophys. J. 2008, 95, 4790. (14) Gross, R.; Wolf, M. M. N.; Schumann, C.; Friedman, N.; Sheves, M.; Li, L.; Engelhard, M.; Trentmann, O.; Neuhaus, H. E.; Diller, R. J. Am. Chem. Soc. 2009, 131, 14868. (15) Herbst, J.; Heyne, K.; Diller, R. Science 2002, 297, 822. (16) Schumann, C.; Gross, R.; Wolf, M. M. N.; Diller, R.; Michael, N.; Lamparter, T. Biophys. J. 2008, 94, 3189. (17) Alexandre, M. T. A.; Domratcheva, T.; Bonetti, C.; van Wilderen, L. J. G. W.; van Grondelle, R.; Groot, M. L.; Hellingwerf, K. J.; Kennis, J. T. M. Biophys. J. 2009, 97, 227. (18) Kondo, M.; Nappa, J.; Ronayne, K. L.; Stelling, A. L.; Tonge, P. J.; Meech, S. R. J. Phys. Chem. B 2006, 110, 20107. (19) Wolf, M. M. N.; Schumann, C.; Gross, R.; Domratcheva, T.; Diller, R. J. Phys. Chem. B 2008, 112, 13424. (20) Nishina, Y.; Sato, K.; Setoyama, C.; Tamaoki, H.; Miura, R.; Shiga, K. J. Biochem. (Tokyo, Jpn.) 2007, 142, 265. (21) Wolf, R.; Reiff, F.; Wittmann, R.; Butzke, J. A process is provided for preparing riboflavin from D-glucose in good yield and 7627

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