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Fluorescence Line Narrowing Studies on Isolated Chlorophyll Molecules Alison Telfer,‡ Andrew A. Pascal,§ Luc Bordes,§ James Barber,‡ and Bruno Robert*,§ DiVision of Molecular Biosciences, Biochemistry Building, Imperial College London, South Kensington Campus, London SW7 2AZ, U.K., and Institut de Biologie et de Technologies de Saclay, CEA, and URA 2096, CNRS, CEA-Saclay, 91191 Gif-sur-YVette, France ReceiVed: August 5, 2009; ReVised Manuscript ReceiVed: December 24, 2009
Fluorescence line-narrowing and resonance Raman properties of various chlorophylls have been measured in organic solvents. Resonance Raman spectroscopy is already a well-established method for the study of photochemical reactions in the various pigment-protein complexes involved in photosynthesis, while fluorescence line-narrowing is still an emerging technique for such systems. Interpretation of these vibrational spectra requires accurate comparative data on the pure isolated pigments. By comparing three different chlorophylls, a, b, and d, which have different substituents on the porphyrin ring, the various spectral lines associated with vinyl and formyl groups on the X and Y electronic axes could be distinguished. The difference between five- and six-coordination of the central Mg atom in FT-Raman spectra was determined by varying the organic solvent used. These chlorophylls are important in photosynthesis: all three in light-harvesting and energy transfer and, in the case of a and d, also in electron transfer. The assignment of spectral bands which we provide here, along with the description of their behavior with respect to the conformation and state of interaction of the pigment molecule, constitutes an essential step if these vibrational techniques are to be exploited to their full potential. Introduction During the photosynthetic process, chlorophyll molecules play multiple roles. In antenna proteins, they trap photons and transfer singlet excitation energy toward reaction center proteins. In the latter they are involved both in charge separation and in the first steps of electron transfer.1 Optimization of each of these functions requires precise tuning of the physicochemical properties of these molecules, in particular their lowest singlet excitedstate energy and their redox potential. Understanding the equilibration of excitation energy within the complex lightharvesting proteins of higher plants and algae, which often bind more than 10 chlorophyll (Chl) cofactors each, as well as the subsequent excitation transfers between these proteins, involves revealing which Chl(s) has the lowest energy singlet excited state. Resonance Raman (RR) spectroscopy constitutes a very effective tool in studying these physicochemical parameters and in assessing their sensitivity to local interactions. It yields specific information on the structure, conformation, and configuration of the molecule studied, via direct measurement of its vibrational energy levels. It has been shown to give selective structural information on BChl, both in the isolated state and when bound to proteins.2,3 For example, the frequencies of a number of RR modes associated with the chlorophyll macrocycle are linearly related to the molecular core size.4 They can therefore be used to assess the coordination state of the central Mg atom5 and can also detect distortions of the macrocycle even to well below 1 Å.6 Modes of conjugated carbonyl groups are sensitive to H-bonds these groups are involved in and to the electrostatic properties of their environment.2 All these parameters may potentially influence the functional properties of * To whom correspondence should be addressed. Tel.: 33-169089015. Fax: 33-169084389. E-mail:
[email protected]. ‡ Imperial College London. § CEA-Saclay.
protein cofactors, and this has often been shown to be the case for bacteriochlorophylls. Indeed, reasonable progress has been made over the past decade in characterizing the mechanisms used by proteins to tune the absorption7,8 and redox9,10 properties of bound Bchl molecules, mediated by both H-bonding7,9 and electrostatics.8,10 The use of RR spectroscopy has been central to these investigations. These studies were relatively straightforward in photosynthetic bacteria, where the number of unique proteins is small and each one only binds a small number of pigments (e.g., 2-3 Bchls per Rβ dimer in LH-type proteins). In addition, producing site-directed mutants for these proteins is routine. This is not the case for chlorophyll-containing organisms, which are both more complex and less readily manipulated. As a result RR is not selective enough to distinguish chlorophyll molecules on the basis of their difference in absorption alone, except in a limited number of cases,11,12 so the equivalent knowledge for protein-bound chlorophylls remains vague. The resonance phenomenon in RR is derived from the coupling of vibrational modes with an electronic transition of the molecule studied. This provides an additional level of selectivity, as only those modes coupled with this transition will be observed. Thus in the case of chlorophylls, wavelengths which correspond to transitions associated with the molecular X-axis of the molecule (i.e., the BX and QX absorption bands) will excite Raman bands for modes vibrating along the X-axis, while modes associated with the Y-axis are seen in BY and QY excitation. An alternative vibrational spectroscopy which also uses the resonance phenomenon is fluorescence line-narrowing (FLN). In contrast to RR spectroscopy, the resonance effect in FLN relies on the fluorescence properties of the emitting molecules. Briefly, the vibrational modes of molecules may be observed when fluorescence is measured with high resolution at very low temperature (below 15 K). In this case, as Chl fluorescence is only observed from the QY transition, only those
10.1021/jp907537a 2010 American Chemical Society Published on Web 01/22/2010
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modes vibrating along the Y-axis can be observed in FLN. The application of FLN for analyzing the vibrational modes of chlorophyll-like molecules was first demonstrated over 30 years ago.13,14 However, this pioneering work was not followed by relevant biological applications until FLN was used, more than 10 years later in the biophysics group of the Free University of Amsterdam,15,16 to obtain the vibrational properties of the terminal emitter(s) of the major light-harvesting protein of higher plants (LHCII). Since then, it has been successfully applied to other light-harvesting chlorophyll-proteins17-19 as well as to the photosystem II reaction center.20 In a sense FLN is even more selective than RR spectroscopy, as it specifically yields information only on the emitter chlorophylls present in these complex pigment-proteins. The information obtained is thus more limited but more precise and relates to the molecular structure of the Chl molecule(s) possessing the lowest energy first singlet excited state.21 However, the information content of chlorophyll FLN spectra has not been fully exploited up to now. This is mainly due to the fact that, in contrast to RR spectroscopy, only a very few reports of the FLN of isolated chlorophyll molecules have been published.14,17,22 This often makes the identification of the vibrational bands in FLN spectra difficult, as attributing these bands usually requires the analysis of the spectra of different model systems (different chemical species and/or different environments). In the case of chlorophylls, such an analysis has only been reported for excitation FLN spectra.23 These yield information on the vibrational properties of the first singlet excited state of the Chl molecules, rather than on their ground state. Due to the extensive excitation energy exchange which occurs between the pigments of light-harvesting proteins, excitation spectroscopy cannot be used on photosynthetic systems - such ultrafast transfers generally prevent the observation of vibrational narrowing, as it induces a blurring of the very fine structures in FLN spectra.24 In the present work we report the FLN spectra of chlorophylls a, b, and d and compare the observed frequencies with those obtained under the same experimental conditions using RR spectroscopy. Most of the spectra have been measured in tetrahydrofuran (THF), a solvent which provides two axial ligands to the central Mg atom of Chl (so that it is sixcoordinated). In some cases spectra were measured in diethyl ether (DEE), which gives only one external Mg ligand at room temperature (i.e., five-coordination). We determine which bands will be most useful for describing the molecular structure of these chlorophylls in vivo. This will broaden the application of chlorophyll FLN spectroscopy to biological systems.
(Coherent, Palo Alto, CA). FLN spectra were recorded with 90° signal collection using a two-stage monochromator (U1000, Jobin-Yvon, Longjumeau, France), equipped with 600 groove/ mm gratings and a front-illuminated, deep-depleted CCD detector (Jobin-Yvon, Longjumeau, France). Typically, less than 1 µW reached the sample during the recording of the spectra. To avoid spurious hole-burning phenomena, which would result in a decrease in the narrow vibrational bands after the first seconds of illumination, the samples were heated to 50 K between individual FLN measurements (which were recorded at 4 K). The spectra shown here were concatenated from ca. 400 cm-1 windows, and the exposure time for recording each window was typically 1-2 s. Each of the displayed spectra thus corresponds to about 8 s of illumination of the sample (the laser beam is blocked by a shutter between measurements). Fourier transform (FT)-Raman spectra were recorded at room temperature using 1064 nm excitation from a YAG laser, as already described.26 The resonance Raman spectrum of Chl d was obtained at 77 K using 441.6 nm excitation from a He-Cd laser (Liconix, Santa Clara, CA), as described previously.7
Methods
Results
Chlorophylls a and b were obtained from Sigma-Aldrich; chlorophyll d was extracted from cells of the predominantly chlorophyll d-containing cyanobacterium, Acaryochloris marina (chlorophyll d g 95% of total chlorophyll), essentially as described by Di Valentin et al.25 Pigment extracts were dried down under a stream of oxygen-free nitrogen or argon and stored in the dark at -20 °C. For FLN measurements, a drop of highly concentrated sample (in THF unless otherwise stated) was deposited on a glass slide (2-3 µL, OD > 10) and frozen in liquid nitrogen prior to insertion in a helium-flow cryostat (Air Liquide, Sassenage, France). This protocol results in large FLN signals, but makes the recording of the spectral region below 650 cm-1 difficult due to the intense light scattering at low frequencies. Excitation in the 640-680 nm range was provided by a dye laser (Spectra Physics, Mountain View, CA) pumped by a 24 W Sabre laser
Figure 1 shows the molecular structure of chlorophylls a, b, and d. The differences in structure between them are as follows: (i) at position 3 (IUPAC numbering) there is a vinyl group in Chls a and b whereas it is a formyl group in Chl d, and (ii) at position 7 in Chl b there is a formyl group in place of the methyl for Chls a and d. In Figure 2 we present high-resolution fluorescence spectra of Chl b in THF at 77 and 4 K, using excitation at 648 nm. Lowering the temperature to 4 K induces a dramatic change in the spectrum, due to the narrowing of a large number of vibrational transitions which appear above the fluorescence envelope. This is the narrowing phenomenon in FLN. We have measured RR and FLN spectra of all three Chls in THF (see Figures 3-5). The frequencies of the FLN bands match remarkably closely with those observed in Raman spectra (see Figures 3 and 5 and Table 1), although, as expected, the
Figure 1. Molecular structure of Chls a, b, and d. The X and Y electronic axes are indicated by dashed lines.
Isolated Chlorophyll Molecules
Figure 2. Effect of temperature on the fluorescence spectrum of Chl b in THF: 77 (lower trace) and 4 K (upper trace).
Figure 3. Comparison of the 4 K FLN and room-temperature FT-RR spectra of Chl b in THF (upper and lower traces, respectively). The FLN spectrum was corrected for the sloping baseline (due to underlying fluorescence) in order to ease comparison of the various modes with those in the FT-RR spectrum. Other details as described in Methods. V, F, and K: contributions of the vinyl, formyl, and keto nuclear coordinates, respectively.
intensities of the observed bands are different for spectra obtained by the two methods. It clearly appears that modes conjugated with the X electronic axis of the molecule are much less active in FLN spectra than in Raman. This is best observed for the vinyl stretching mode at 1624 cm-1 and the formyl carbonyl stretching mode at 1656 cm-1 (V, F in Figures 3-5). These modes involve a limited number of nuclear coordinates and are mainly conjugated with the X electronic axis.27 Figure 4 shows a comparison between the FLN spectra of Chl b (obtained at 651 nm) and Chl a (at 677 nm). As already reported from resonance Raman studies,28 the vibrational spectra of these two pigments are quite different, due to the presence of a formyl carbonyl on the Chl b molecule (Figure 1). Indeed it has been shown, using chemically modified pigments, that the presence (or removal) of such a chemical group induces dramatic changes in the vibrational spectra of both bacteriochlorophyll and chlorophyll-like molecules.29 However, it should be noted that, contrary to resonance Raman spectra, the analysis of the higher frequencies of these FLN spectra, where the stretching modes of the conjugated carbonyl groups contribute,
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Figure 4. FLN spectra (obtained at 4 K) of Chls b (upper trace) and a (lower trace) in THF. Spectra were not baseline corrected. V, vinyl contributions. Other bands referred to in the text are indicated by their respective numbers or by asterisks.
Figure 5. Comparison of vibrational spectra of Chl d in THF - FLN (upper trace; obtained at 4 K) and RR (lower trace; obtained at 77 K, excitation 441.6 nm). F and K: formyl and keto contributions, respectively. Other modes referred to in the text are indicated by the appropriate number.
cannot be used to distinguish between Chls a and b, as the FLN band arising from the stretching mode of the formyl carbonyl of Chl b is not observed in these spectra. Figure 5 displays the FLN spectrum of isolated Chl d, excited at 707 nm, as compared to the RR spectrum of this molecule obtained at 441.6 nm. Again, the presence of an additional conjugated formyl carbonyl on this molecule as compared to Chls a and b, in this case at position 3 in place of a vinyl group (see Figure 1), induces dramatic changes in both the frequencies and intensities of the vibrational bands in the Chl d spectra (compare Figures 4 and 5). All the observed frequencies in Figures 3-5 are reported in Table 1, along with those for room temperature FT-Raman spectra of Chl a in both THF and DEE. These solvents provide two and one axial ligand(s), respectively, to the central Mg of this molecule under these conditions. Discussion In this work, we report FLN spectra of three isolated forms of chlorophyll: a, b, and d in THF, a solvent which provides
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TABLE 1: Vibrational Bands of Chls a, b, and da Chl a, THF FLN FT-RR
Chl b, THF
Chl a, DEE Chl a* FT-RR FLN FT-RR FLN Raman FLN
693 750 789
744
742
785
841
1045 1071
1102 1120 1145
1121 1140
R7 1181 1204 1224 1237 1264 / 1289 1308 V 1325 1348
1185 1206 1223 1234 1263 1291 1306 1326 1347
1384 R6 1430 R5 1485
1387 1398 1435 1485
R4 1526 R3 1549 R2 R1 1594 V F K
/
1688
698 738 745 756 777 793
757
835
916 926 984 1030 R8 1048 1071
918 922 986
Chl d, THF
911 985
919 942 976 1041
1069
1091
1114 1141 1146 1184 1203 1224
1118
922 976 1005
690 720 737 750
798 817 839 897 908 924 983 1006
794 820 858
1154 1181
1155 1181
1225 1233 1266 1291
1224
1119 1136 1148 1170 1205 1234
1266 1308
1263 1289
1322 1350
1323 1351
1388
1388
1391
1436 1490
1429 1474
1437 1474
1528 1550 1580 1597
1533 1551 1584 1607
1515 1546 1562 1592 1601
1535 1563 1598
1625
1625
1689
1701
1262 1287 1304 1328 1345
1699
1123
688 723 744 766
1627 1663 1701
1346 1351 1377 1395
913 983 1004 1019 1074
1136 1164 1224 1235 1263 1295 1345
1586
1377 1388 1398 1422 1482 1495 1529 1545 1574 1590
1656 1693
1662 1692
1477 1518 1540
687 715 735 760 780 802 843 877 902 912 921 981 1004 1035 1064 1075 1122 1132 1168 1192 1224 1232 1250 1274 1315 1325 1351 1376 1403 1415 1426 1514 1535
1657
a
Data on FLN of Chl a* in triethylamine is taken from ref 23. Bands (potentially) sensitive to molecular core size are indicated by R1-8 and / (see text for details). V, F, K: vinyl, formyl and keto contributions.
two axial ligands to the central Mg of these molecules (so it is six-coordinated). It is not easy to obtain FLN spectra of chlorophyll with only one axial ligand, as the narrowing phenomenon can only be observed at very low temperatures (see Figure 2). Lowering the temperature of samples of fivecoordinated Chl in solvents induces, in most cases, a shift to a mixture of five- and six-coordinated species, or even to sixcoordination only (this is the case, for instance, for diethyl ether17,30). In the absence of studies of actual chlorophyll models by FLN, it is thus hard to predict the information content of the observed bands in the FLN spectra. A parallel can be made with the bands observed in resonance Raman spectra (which can be measured at higher temperatures). However, RR spectra of sufficient quality are difficult to obtain for chlorophyll molecules in resonance with their QY electronic transition due to their intrinsic fluorescence in this region. RR spectra are more
easily obtained in Soret excitation conditions,11,12,28,31 but comparing the sensitivity of bands obtained in resonance with different electronic transitions can be problematic (note that Chl FLN represents resonance with the QY transition). Indeed, it is known that, in different resonance conditions, bands may contribute at the same position in the spectra but arise from slightly different vibrational modes and thus are not sensitive to the same molecular parameters.17 FT-Raman spectra of Chl molecules excited at 1064 nm, in preresonance with the Chl QY transition, should primarily contain similar contributions to those seen in FLN spectra. As mentioned above, this is indeed the case: the same bands are seen in both spectra, although with different intensities. It is therefore of use to determine the sensitivity of bands in FT-Raman spectra to the molecular conformation of Chl molecules. This analysis can then be used as an indirect indicator of the sensitivity of the equivalent bands in FLN spectra. In the mid-1980s Fujiwara and Tasumi studied a number of metal-substituted chlorophylls using resonance Raman spectroscopy under Soret excitation conditions and described a series of Raman bands which give direct information on the molecular conformation of these molecules.32,33 They showed that the frequency of no less than eight modes contributing between 1000 and 1600 cm-1 is linearly dependent on the core size of the molecule, i.e. the average distance between the central metal atom and the pyrrolic nitrogens. In the case of Chl a, these modes came at 1048, 1186, 1438, 1495, 1530, 1558, 1588, and 1614 cm-1 (termed R8 to R1, respectively33). As discussed by Nave¨ke et al.,5 the modes sensitive to the core size of these molecules are also dependent on the coordination state of their central Mg atom, as a change in coordination generally induces a change in the Chl molecular geometry. It is thus possible to verify the presence of the R1-8 modes in the spectra displayed in this work by comparing FT-Raman spectra of five- and sixcoordinated Chl a (measured in DEE and THF, respectively). At the frequencies of the R8 mode, only extremely weak bands contribute to the FT-RR spectra obtained in preresonance with the Chl a QY transition. At the frequencies where R3, R6, and R7 contribute, the bands observed here show no dependence on the coordination state of Chl a (see Table 1). This indicates that although the frequency of these bands matches with those observed in Soret excitation, the modes they arise from involve different sets of nuclear coordinates. By contrast, the bands which contribute at the frequencies of the R1, R2, R4, and R5 modes display a clear dependence on the state of coordination of the central Mg of Chl a, which matches with that expected from Fujiwara’s work.32,33 These bands thus probably arise from the same vibrational modes in Soret and QY excitation. Finally, in preresonance conditions with the QY electronic transition, two supplementary bands located at ∼1120 and ∼1290 cm-1 also display a clear sensitivity to the Mg coordination state (indicated in Table 1 and Figure 4 by asterisks). Remarkably these bands downshift when this atom goes from six- to five-coordination (the other four are up-shifted). As stated above, the frequency of the bands observed in FLN and FT-Raman spectra of Chl a show remarkable similarity. It is thus likely that the modes described as sensitive to molecular conformation of this molecule in FT-Raman spectra are conserved in FLN spectra. Care must be taken, however, as the two methods rely on different selection rules. The six modes sensitive to conformation in Chl a FT-Raman spectra vibrate at 1597, 1580, 1528, 1485, 1291, and 1121 cm-1 for sixcoordinated Chl a (in THF) and at 1607, 1584, 1533, 1490, 1287, and 1114 cm-1 for Chl a molecules with a five-
Isolated Chlorophyll Molecules coordinated central Mg atom (in DEE; see Table 1). In FLN spectra of six-coordinated Chl a (Figure 4, lower trace), these modes are observed at 1594, (∼1580), 1526, 1485, 1289, and 1120 cm-1 (the R2 mode at 1580 cm-1 is only a very weak shoulder). To aid in their identification, these modes are labeled in Figure 4 by their corresponding numbers or, for the two newly described modes, by asterisks. In FLN spectra of LHCII complexes, where most of the Chls a are five-coordinated, these modes contribute at 1612, (∼1580), 1537, 1487, 1286, and 1110 cm-1.16 It thus seems quite clear that these five bands (excluding the weak R2 mode) in the FLN spectra are fingerprints of the coordination state of Chl a in much the same way as their counterparts in FT-Raman spectra, at least through their sensitivity to the molecular core size of this molecule. Finally these spectra may allow a direct comparison between the vibrational modes of the ground and first excited states of Chl a. Table 1 compares the band frequencies of the FLN spectra reported in this work with those in FLN excitation spectra (Chl a*) obtained by Renge et al.23 The downshift of the stretching mode of the keto carbonyl of Chl a between these spectra is consistent with that reported by time-resolved FTIR spectroscopy.34 Altogether the resonance Raman spectrum of Chl d displayed in this paper is very similar to those reported by Chen et al.,35 although there are some differences. Most of these differences may be explained by the different experimental conditions in which the spectra were obtained. Chen et al.35 prepared their sample by drying an acetonitrile solution of Chl d, which is expected to induce self-association. As discussed below, the band frequencies reported by Chen et al. are consistent with Chl d having only one axial ligand (with a five-coordinated central Mg), a situation difficult to observe in solution but which generally appears upon self-association of Chl molecules. In addition the keto carbonyl contribution indicates that this grouping is H-bonded (see below), again consistent with selfassociation. As the spectra we report here are for Chl d in THF, i.e., with a six-coordinated central Mg atom, comparing them with Chen’s spectra35 may again help in determining which vibrational bands are sensitive to the core size of this molecule. It clearly appears that at least four of the eight modes defined by Fujiwara and Tasumi33 are present in Chl d spectra and display different frequencies according to the coordination state of the central Mg. These modes are R1, R3, R4, and R5sobserved at 1590, 1545, 1529, and 1482 cm-1 in this work (indicated in Figure 5) and at 1610, 1554, 1533, and 1493 cm-1 in Chen’s spectra.35 Just as for Chl a, these Chl d modes are most likely to be sensitive to the Mg coordination state through their sensitivity to the molecular core size. We also observe that some bands reported in this work are missing in Chen’s spectra,35 such as the one at 1574 cm-1. This is probably due to the fact that the spectra have not been obtained with the same resolution, which often makes the observation of weak bands located at the foot of very intense ones difficult to observe. Finally, we observe here a band at 1692 cm-1, which arises from the stretching mode of the keto carbonyl group of Chl d when it is free from intermolecular interactions. This band is only extremely weak in Chen’s spectra35 because Chl d self-associates via H-bonding of this group, inducing a down-shift in its stretching frequency. Indeed a much larger contribution is observed in Chen’s spectra as a shoulder on the high-frequency side of the 1660 cm-1 band,35 arising from the stretching mode of the keto group when it is involved in an intermolecular H-bond.
J. Phys. Chem. B, Vol. 114, No. 6, 2010 2259 As observed in resonance Raman, the introduction of additional carbonyl groups into the chlorophyll structure induces dramatic perturbations of the vibrational properties of the molecule. First additional bands arising from the stretching modes of these carbonyls can be observed (around 1660 cm-1 for the formyls of both Chl b and Chl d). It is of note that, in FLN spectra, the band arising from the stretching mode of the formyl carbonyl of Chl b can barely be observed, as it is coupled almost entirely with the X electronic axis of the molecule, while a clear band is present at that frequency in FT-Raman spectra of this molecule (Figure 3). By contrast, the stretching mode of the formyl carbonyl of Chl d results in a very clear band in FLN spectra (Figure 5), as should be expected from the position of this group on the molecule (along the Y axis; see Figure 1). In the middle-frequency range, the bands contributing to the Chl d FLN spectrum display very similar frequencies to those in resonance Raman spectra (Figure 5, Table 1). However, the RR band at 1545 cm-1 is shifted to 1540 cm-1 in FLN spectra. By analogy with the FLN spectra of Chl a, we hypothesize that this band arises from slightly different vibrational modes, and it thus may not be sensitive to the conformation of the Chl d molecule. There is a band at 1289 cm-1, essentially the same frequency as the ∼1290 cm-1 band in Chl a spectra which is sensitive to molecular conformation, but, in the absence of comparison between five- and six-coordinated Chl d, formal conclusions about the sensitivity of this band cannot be made. In the 600-1000 cm-1 region (Figure 5), the frequency of the observed bands in FLN and RR are also similar, but their intensities are totally different. Finally, the spectra of Chl d reported in this paper may be compared to the spectra of acetyl-chlorophyll a reported by Feiler et al.29 In the latter work, modes mainly involving the nuclear coordinates of the vinyl group of Chl a were proposed to contribute to the spectra at 1625 and 1327 cm-1. It is of note that both these bands are absent in the RR spectrum of Chl d reported here, which lacks the vinyl group (Figure 1), whereas they are clearly present for Chls a and b (Figures 3 and 4; see Table 1). The RR spectrum of Chl d above 1260 cm-1 is remarkably similar to that of 2-acetyl Chl a. However, both the positions and the intensities of the bands observed in the lower frequency range clearly differ between these two molecules, indicating that there are different effects on the conjugated macrocycle of these slightly different groupings. Here we have compared FLN and resonance Raman spectra of various chlorophylls in their ground state, and used their chemical differences to assign specific modes to specific groupings on the molecule. By observing the effects on the spectra of using different solvents we also describe the influence of molecular conformation on these band frequencies (through the coordination state of the central Mg). These attributions should prove fundamental to the application of these vibrational techniques to the investigation of chlorophyll functions in biological materials. Acknowledgment. We are very grateful to Mr. Alan Scott at Queen Mary, London University for assistance in carrying out HPLC during the purification of Chl d. This work was supported by the French National Agency for Research (ANR), by projects MASTRITT and CAROPROTECT, by the U.K. Biotechnology and Biological Sciences Research Council and by the EU Marie Curie program (FP7 International Training Network Harvest). This paper is dedicated to Prof. Dr. Achim Trebst on the occasion of his 80th birthday.
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References and Notes (1) Blankenship, R. E. Molecular Mechanisms of Photosynthesis; Blackwell Science: Oxford, U.K., 2002. (2) Lapouge, K.; Nave¨ke, A.; Sturgis, J.; Hartwich, G.; Renaud, D.; Simonin, I.; Lutz, M.; Scheer, H.; Robert, B. J. Raman Spectrosc. 1998, 29, 977–981. (3) Gall, A.; Robert, B. Biochemistry 1999, 38, 5185–5190. (4) Fujiwara, M.; Tasumi, M. Nippon Kagaku Kaishi 1988, 566–568. (5) Nave¨ke, A.; Lapouge, K.; Sturgis, J.; Hartwich, G.; Simonin, I.; Scheer, H.; Robert, B. J. Raman Spectrosc. 1997, 28, 599–604. (6) Lapouge, K.; Nave¨ke, A.; Gall, A.; Ivancich, A.; Seguin, J.; Scheer, H.; Sturgis, J.; Mattioli, T.; Robert, B. Biochemistry 1999, 38, 11115–11121. (7) Sturgis, J.; Robert, B. J. Phys. Chem. B 1997, 101, 7227–7231. (8) Gall, A.; Fowler, G.; Hunter, C.; Robert, B. Biochemistry 1997, 36, 16282–16287. (9) Mattioli, T. A.; Lin, X.; Allen, J. P.; Williams, J. C. Biochemistry 1995, 34, 6142–6152. (10) Spiedel, D.; Jones, M. R.; Robert, B. FEBS Lett. 2002, 527, 171– 175. (11) Pascal, A.; Wacker, U.; Irrgang, K.; Horton, P.; Renger, G.; Robert, B. J. Biol. Chem. 2000, 275, 22031–22036. (12) Pascal, A.; Caffarri, S.; Croce, R.; Sandona, D.; Bassi, R.; Robert, B. Biochemistry 2002, 41, 2305–2310. (13) Fu¨nfschilling, J.; Williams, D. F. Photochem. Photobiol. 1975, 22, 151–152. (14) Rebane, K.; Avarmaa, R. Chem. Phys. 1982, 68, 191–200. (15) Kwa, S. L. S.; Vo¨lker, S.; Tilly, N. T.; van Grondelle, R.; Dekker, J. P. Photochem. Photobiol. 1994, 59, 219–228. (16) Peterman, E.; Pullerits, T.; van Grondelle, R.; van Amerongen, H. J. Phys. Chem. B 1997, 101, 4448–4457. (17) Pascal, A.; Peterman, E.; Gradinaru, C.; van Amerongen, H.; van Grondelle, R.; Robert, B. J. Phys. Chem. B 2000, 104, 9317–9321. (18) Kleima, F.; Wendling, M.; Hofmann, E.; Peterman, E.; van Grondelle, R.; van Amerongen, H. Biochemistry 2000, 39, 5184–5195.
Telfer et al. (19) Ratsep, M.; Pieper, J.; Irrgang, K.; Freiberg, A. J. Phys. Chem. B 2008, 112, 110–118. (20) Peterman, E.; van Amerongen, H.; van Grondelle, R.; Dekker, J. Proc. Natl Acad. Sci. U.S.A. 1998, 95, 6128–6133. (21) Avarmaa, R.; Rebane, K. Spectrochim. Acta A: Mol. Biomol. Spectrosc. 1985, 41, 1365–1380. (22) Ratsep, M.; Linnanto, J.; Freiberg, A. J. Chem. Phys. 2009, 130, 194501–194511. (23) Renge, I.; Mauring, K.; Sarv, P.; Avarmaa, R. J. Phys. Chem. 1986, 90, 6611–6616. (24) Avarmaa, R.; Jaaniso, R.; Mauring, K.; Renge, I.; Tamkivi, R. Mol. Phys. 1986, 57, 605–621. (25) Di Valentin, M.; Ceola, S.; Agostini, G.; Telfer, A.; Barber, J.; Bohles, F.; Santabarbara, S.; Carbonera, D. Mol. Phys. 2007, 105, 2109– 2117. (26) Mattioli, T.; Hofmann, A.; Sockalingum, D.; Schrader, B.; Robert, B.; Lutz, M. Spectrochim. Acta A: Mol. Biomol. Spectrosc. 1993, 49, 785– 799. (27) Feiler, U.; Albouy, D.; Lutz, M.; Robert, B. Photosynth. Res. 1994, 41, 175–180. (28) Pascal, A.; Caron, L.; Rousseau, B.; Lapouge, K.; Duval, J.; Robert, B. Biochemistry 1998, 37, 2450–2457. (29) Feiler, U.; Mattioli, T.; Katheder, I.; Scheer, H.; Lutz, M.; Robert, B. J. Raman Spectrosc. 1994, 25, 365–370. (30) Robert, B.; Lutz, M. In Spectroscopy of Biological Molecules; Alix A. J. P., Ed.; John Wiley & Sons: Chichester, U.K., and Reims, France, 1983; pp 338-41. (31) Germano, M.; Pascal, A.; Shkuropatov, A.; Robert, B.; Hoff, A.; van Gorkom, H. Biochemistry 2002, 41, 11449–11455. (32) Fujiwara, M.; Tasumi, M. J. Phys. Chem. 1986, 90, 250–255. (33) Fujiwara, M.; Tasumi, M. J. Phys. Chem. 1986, 90, 5646–5650. (34) Groot, M.; Breton, J.; van Wilderen, L.; Dekker, J.; van Grondelle, R. J. Phys. Chem. B 2004, 108, 8001–8006. (35) Chen, M.; Zeng, H.; Larkum, A.; Cai, Z. Spectrochim. Acta A: Mol. Biomol. Spectrosc. 2004, 60, 527–534.
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