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Feb 4, 2009 - (1) Violaxanthin is a xanthophyll relatively weakly bound to the ... of isolated LHCII results in energetic uncoupling of violaxanthin f...
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J. Phys. Chem. B 2009, 113, 2506–2512

Light-induced Change of Configuration of the LHCII-Bound Xanthophyll (Tentatively Assigned to Violaxanthin): A Resonance Raman Study Wiesław I. Gruszecki,*,† Małgorzata Gospodarek,‡ Wojciech Grudzin´ski,† Radosław Mazur,§ Katarzyna Gieczewska,§,| and Maciej Garstka§ Department of Biophysics, Institute of Physics, Maria Curie-Skłodowska UniVersity, 20-031 Lublin, Poland, Institute of Physics, Technical UniVersity of Lublin, Lublin, Poland, Department of Metabolic Regulation and Department of Plant Anatomy and Cytology, Faculty of Biology, UniVersity of Warsaw, Miecznikowa 1, 02-096 Warsaw, Poland ReceiVed: NoVember 19, 2008; ReVised Manuscript ReceiVed: December 31, 2008

Raman scattering spectra of light-harvesting complex LHCII isolated from spinach were recorded with an argon laser, tuned to excite the most red-absorbing LHCII-bound xanthophylls (514.5 nm). The intensity of the ν4 band (at ca. 950 cm-1) corresponding to the out-of-plane wagging modes of the C-H groups in the resonance Raman spectra of carotenoids appears to be inversely dependent on the probing laser power density. This observation can be interpreted in terms of excitation-induced change of configuration of the proteinbound xanthophyll owing to the fact that the intensity of this particular band is diagnostic of a chromophore twisting resulting from its binding to the protein environment. The comparison of the shape of the ν4 band of a xanthophyll involved in the light-induced spectral changes with the shape of the ν4 band of the xanthophylls present in LHCII, reported in the literature, lets us conclude that, most probably, violaxanthin is a pigment that undergoes light-driven changes of molecular configuration but also the involvement of lutein may not be excluded. Possible physical mechanisms responsible for the configuration changes and physiological importance of the effect observed are discussed. 1. Introduction Light-harvesting pigment-protein complex LHCII, a major photosynthetic antenna, is a trimer composed of three Lhcb proteins (Lhcb1-3). Each monomer comprises eight molecules of chlorophyll a, six molecules of chlorophyll b, and four molecules of xanthophyll pigments: two luteins, one neoxanthin, and one violaxanthin.1 Violaxanthin is a xanthophyll relatively weakly bound to the protein environment of LHCII and therefore its actual stoichiometry is strongly dependent on both the complex isolation procedure and the treatment.2 It has been reported that illumination of isolated LHCII results in energetic uncoupling of violaxanthin from the chlorophyll pool, manifested by alteration of the chlorophyll a fluorescence excitation spectrum.3,4 It has been suggested that trans-cis isomerization of violaxanthin may be involved in this process.5,6 In this report we present results of the experiments carried out with the application of the resonance Raman (RR) scattering technique, which can be interpreted in terms of light-induced change of the configuration of the LHCII-bound xanthophyll violaxanthin. RR scattering has been demonstrated to be a powerful technique that can be applied to precisely monitor conformational properties of the LHCII-bound xanthophyll pigments and to obtain an insight into molecular dynamics of LHCII.7 Owing to the differences in the energy of the 11Ag-f11Bu+ transition and differences in the dielectric properties of the microenvironment * Corresponding author: E-mail: [email protected]; fax: +4881 537 61 91. † Maria Curie-Skłodowska University. ‡ Technical University of Lublin. § Department of Metabolic Regulation, Faculty of Biology, University of Warsaw. | Department of Plant Anatomy and Cytology, Faculty of Biology, University of Warsaw.

of the LHCII-bound xanthophylls, the positions of the 0-0 vibrational transitions in their in situ absorption spectra vary considerably. This band has been localized at 488 nm for neoxanthin, at 492 nm for violaxanthin, and at 489 and 495 nm for two lutein molecules differing distinctly in spectroscopic properties.8 Recent analysis of the RR spectra of intact leaves and of isolated thylakoid membranes and LHCII, recorded with the 488 nm laser excitation, has revealed a light-induced conformational rearrangement of the protein-bound xanthophyll neoxanthin.9 In the present work we apply the 514.5 nm Argon laser line to selectively excite the LHCII-bound xanthophylls absorbing light in the long-wavelength edge of the Soret band. The comparison of the RR spectra recorded with different probing light intensities reveals an operation of a molecular mechanism that consists in light-dependent configuration change of the LHCII-bound xanthophyll(s). 2. Materials and Methods 2.1. LHCII and Violaxanthin Isolation. The largest lightharvesting antenna complex, LHCII, was isolated from fresh spinach leaves according to a slightly modified procedure of Krupa and co-workers.10 To ensure high purity of the preparation, the final step of the protein precipitation in 20 mM MgCl2 and 100 mM KCl has been additionally repeated for the third time and followed by a 40 min centrifugation at 30 000g on a sucrose gradient. The preparation was suspended in a Tricine buffer (20 mM, pH 7.6) containing 10 mM KCl. Directly before the measurements the LHCII preparation was subjected to centrifugation at 15 000g, and the pellet containing aggregated protein was suspended with Tricine buffer (20 mM, pH 7.6) containing 10 mM KCl. The preparation presented typical of LHCII electronic absorption, fluorescence, and FTIR spectra.11-14 The purity of the preparation has been further tested by means

10.1021/jp8101755 CCC: $40.75  2009 American Chemical Society Published on Web 02/04/2009

Light-harvesting Pigment-Protein Complex LHCII

Figure 1. Electrophoretical analysis of LHCII preparation. Panel a: native electrophoresis pattern of LHCII:LHCIIn, oligomeric; LHCII3, trimeric; LHCII, monomeric forms. Panel b: immunodetection of major Lhcb proteins.

of electrophoresis, mass spectrometry, and pigments chromatography. The all-trans isomer of violaxanthin was isolated from Viola tricolor blossoms and lutein from spinach leaves and purified chromatographically on a C-30 coated reversed-phase column from YMC GmbH, Germany (length 250 mm, internal diameter 4.6 mm) with the following solvent system: acetonitrile/methanol/water (72:8:3, v/v), which was used as a mobile phase, as described previously in detail.15 Before recording RR spectra, the pigments were transferred to CS2. 2.2. Electrophoresis of LHCII Complex. Native LHCII complex was analyzed by SDS-PAGE electrophoresis followed by immunodetection. Primary antibodies against Lhcb1, Lhcb2, and Lhcb3 proteins (Agrisera) and secondary antibody conjugated with alkaline phosphatase (BioRad) were used. Mildly denaturing electrophoresis was performed as described in detail elsewhere16 with some modifications. 2.3. Mass Spectrometry Measurements. LHCII analysis was performed by reverse-phase liquid chromatography using a Waters 600 HPLC system and poly(styrene-divinylbenzene) copolymer (Polymer Laboratories PLRP/S; 5 µm × 300 Å; 2.1 × 150 mm) stationary phase column. Proteins were eluted by stepped acetonitrile linear gradient, and mass spectra were recorded on a Waters-Micromass ZQ quadrupole mass spectrometer with an electrospray ion source (ESI/MS). Identification of proteins from LHCII complex was performed by comparison of obtained masses with published sequences using PeptideMass software (http://expasy.org/tools/peptide-mass.html). 2.4. Resonance Raman spectroscopy. Raman scattering spectra from the liquid samples placed in the 0.2 mm quartz cuvettes were recorded with the inVia Reflex Raman Microscope from Renishaw (UK) equipped with two holographic ultrahigh precision diffraction grating stages and high sensitivity ultralow noise CCD detector. A 514.5 nm Ar+ laser has been applied to record Raman scattering. Laser power has been tuned in the range 0.08-8 mW. The laser beam diameter was focused either to 3 µm or to 20 µm. The spectra have been accumulated within 10 s integration time. All types of the spectra were recorded at least four times, and the spectral effects reported were found to be reproducible. Resonance Raman scattering spectra were corrected by subtracting a background signal originating from fluorescence. 3. Results and Discussion Examination of fine spectroscopic effects observed in the LHCII protein requires very pure and well-defined preparations. Purity of LHCII preparation was analyzed by gentle detergent treatment followed by mildly denaturing electrophoresis.16 Presented electrophoretic pattern (Figure 1a) reveals green bands exclusively related to oligomeric, trimeric with some monomeric forms of LHCII, and absolute lack of bands assigned to PSII and PSI core complexes and LHCI antenna, usually present in

J. Phys. Chem. B, Vol. 113, No. 8, 2009 2507 crude thylakoid membranes.16,17 High purity of the LHCII preparation was confirmed by immunodetection, which showed dominance of Lhcb1 and Lhcb2 proteins (Figure 1b). Detailed analysis of the LHCII preparation by means of HPLC-ESI/MS (Figure 2) has shown three Lhcb1, two Lhcb2 isoforms, as well as traces of the phosphorylated form of Lhcb1. Although Lhcb3 was not detected by antibodies (Figure 1b), this protein was revealed by mass spectrometry (Figure 2). Quantitative mass spectrometry analysis of LHCII preparation has shown Lhcb1/ Lhcb2/Lhcb3 proteins ratio as 12:6:1. The chlorophyll a to chlorophyll b ratio in the preparation was 1.3, and the xanthophyllsslutein, neoxanthin, and violaxanthinswere present in the ratio of 2.00:1.08:0.51 per protein monomer, respectively, in the samples used for experiments. No zeaxanthin has been detected in the preparation. Application of the 514.5 nm line of the Ar+ laser in RR examination of pure LHCII results in excitation of only two out of four of the protein-bound xanthophylls: the lower wavelength-absorbing lutein (the light absorption maximum of the 0-0 band at 495 nm8) and violaxanthin (the maximum of the 0-0 absorption band at 492 nm8). The 0-0 absorption bands of the other lutein molecule (489 nm) and neoxanthin (488 nm)8 are beyond the resonance. Figure 3 presents the superimposed RR spectra of LHCII, lutein, and violaxanthin normalized at the maximum. Four principal spectral regions can be distinguished in the RR spectra of carotenoid pigments:7 the ν1 band centered in the region of 1525 cm-1, corresponding to the CdC stretching vibrations in the polyene chain; the ν2 band in the region between 1100 and 1250 cm-1, representing the C-C stretching vibrations coupled either to the C-H in-plane bending or to the C-CH3 stretching modes; the ν3 band at ca. 1000 cm-1, corresponding to the CH3 in-plane rocking vibrations; and the ν4 band at ca. 950 cm-1, corresponding to the out-of-plane wagging modes of the C-H groups.18 Owing to the fact that those particular C-H vibrations are neither associated with pronounced polarizability changes nor coupled (for reasons of symmetry) with electronic transitions of the planar conjugated double bond chain of carotenoids, the ν4 band is of relatively low intensity (see Figure 3). On the contrary, the same band is quite well pronounced in infrared absorption vibrational spectroscopy, due to the relatively strong transition dipole moment of this vibration.19-21 As can be seen, lutein and violaxanthin in CS2 are shifted only slightly despite the difference in conjugation length by one double bond. Conjugation length correlates with the position of the ν1 band on wavenumber scale: as the chain becomes progressively longer, ν1 decreases. Such a relationship is somehow affected when conjugation is not strictly linear and extends to the terminal carotenoid rings, for example, the ν1 band of β-carotene is located at higher wavenumbers than in the case of lycopene (conjugation length in both cases is the same, n ) 11).22 Interestingly, the ν4 band of xanthophylls gains intensity upon the pigment binding into the protein environment of the photosynthetic antenna complexes.7 This effect can be seen from the comparison of the spectra presented in Figure 3. Such an effect has been explained in terms of a distortion of the planar xanthophyll configuration in the protein bed, induced by rotations around the C-C bonds.7,23 This distortion results in increased coupling of C-H wagging modes with the polyene vibrations and electronic transitions and consequently results in increased RR spectra intensity.7,23 Similar enhancement accompanying the xanthophyll configuration changes upon binding into the protein environment can be also observed in the case of the ν2 and ν3 bands (Figure 3). Figure 4 presents

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Figure 2. Mass spectroscopy identification of Lhcb proteins in LHCII preparation. Each panel shows mass spectrum for different peaks in HPLC separation and molecular masses, calculated from four independent HPLC runs.

Figure 3. Resonance Raman spectra of isolated lutein, violaxanthin, and LHCII, as indicated. The spectra were normalized in the maximum of the ν1 band. The spectra recorded with the laser power density of 11 µW/µm2.

the normalized RR spectra of LHCII, and Figure 5 presents the ν4 spectral region recorded with different laser light power density. As can be seen, the intensity of the entire ν4 band differs distinctly, despite the fact that the spectra have been normalized at the maximum of the ν1 band. The higher the light intensity, the lower the intensity of the ν4 band. Such a result can be interpreted in terms of light-driven changes of the xanthophylls’ configuration: namely, the liberation from the twisted configuration. This effect is accompanied with the slight increase in overall fluorescence signal recorded as a background of the RR scattering (see Figure 6). The effect of the decrease in the intensity of the ν4 band is not linearly dependent on light

Figure 4. Resonance Raman spectra of LHCII, recorded with different laser power density, as indicated. The spectra were normalized in the maximum of the ν1 band.

intensity (see Figure 7) and represents rather a saturation-type relationship (maximum decrease by ca. 40%). This is consistent with the interpretation that not entire pool of the pigments that are in resonance at 514.5 nm is involved in the light-dependent process observed. Figure 8 presents the comparison of the difference spectrum representing the effect of illumination, calculated on the basis of the spectra presented in Figure 5 with the resonance Raman spectra in the ν4 region, of the xanthophyll pigments of LHCII, based on the literature.24 The spectrum that represents the light effect on LHCII-bound xanthophylls differs distinctly from the spectra of lutein and neoxanthin and closely resembles the spectrum of violaxanthin in LHCII.24 On the basis of such a comparison, one can postulate that the spectral effect

Light-harvesting Pigment-Protein Complex LHCII

Figure 5. Resonance Raman spectra of LHCII presented in the ν4 band spectral region recorded with different laser power density, as indicated. Note that the spectra were normalized in the maximum of the ν1 band (see Figure 4).

Figure 6. Resonance Raman spectra of LHCII, recorded with different laser power density, as indicated. The spectra have not been corrected by subtracting a background originating from the fluorescence signal. The spectra are presented on a wavenumber scale (not calculated as a Raman shift). The fluorescence spectra were normalized so that the relative intensity of the ν1 Raman band, visible on the top of each fluorescence spectrum, equals 1. Owing to such a procedure, the integrated intensities of the fluorescence spectra remain in proportion to the fluorescence quantum yields in different samples (the intensity of the ν1 band in each sample is proportional to light intensity). Such an assumption is valid for low (nondestructive) laser power density values. For higher laser power densities fluorescence signal can be only analyzed qualitatively, as a demonstration of the phenomenon.

observed is related to a light-induced change of the molecular configuration of violaxanthin. On the other hand, contribution to the effect of one lutein molecule that is in resonance at 514.5 nm may not be excluded. Interestingly, violaxanthin present in our preparation at an average concentration 0.5 molecule per LHCII monomer, accounts for approximately 40% intensity in the ν4 band in the RR spectra of the xanthophylls that are in

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Figure 7. Dependence of the integrated intensity of the ν4 band of LHCII (in the range 935-985 cm-1) and of the integrated fluorescence (in the range 16 235-19 286 cm-1) on laser power density of the excitation beam applied. The intensities are presented on an arbitrary unit scale on which the levels corresponding to the lowest light power density are normalized to unity. Before integration, the RR spectra were normalized in the maximum of the ν1 band (as in Figures 3-5). The fluorescence spectra were normalized as described in the legend to Figure 6.

resonance at 514.5 nm. Because the intensity of this band is almost entirely dependent on twisting of the polyene chain and not simply proportional to pigment concentration, such an intense ν4 band suggests a relatively strong deformation of violaxanthin molecule upon binding to the protein environment of LHCII (possibly due to simultaneous interaction with two trimer-forming LHCII polypeptides). The decrease in intensity of the ν4 band, which can be interpreted in terms of relaxation of molecular twisting, is associated with relatively small enhancement of fluorescence quantum yield of LHCII-bound xanthophylls that absorb light at 514.5 nm (see Figure 6 and Figure 7). The recorded fluorescence emission appears in the spectral range typical of the S2 f S0 (1Bu+ f 1Ag-) transition.25,26 The emission bands are rather complex, and even at relatively low fluorescence yield (corresponding to laser power density of 113 µW/µm2) at least 5 Gaussian components are required to reconstitute the emission band (Figure 9), which indicates that, most probably, more than one LHCII-bound xanthophyll is involved in the light emission observed. The bands centered at 16 273, 17 654, and 19 036 cm-1 can be assigned to the 0-3, 0-2, and 0-1 vibronic transitions, respectively, of one pigment, and the second set of components, centered at 17 007 and 18 277 cm-1 can be assigned to the 0-3 and 0-2 vibronic transitions of another xanthophyll. Apparently, the 0-0 transition is beyond the range of our RR spectrometer. It seems difficult to assign those components to particular xanthophylls without falling into speculations. At higher light intensities the fluorescence yield is higher and the band is even more complex. At low probing light intensities the xanthophyll fluorescence quantum yield is very low, which is consistent with very efficient singlet excitation energy transfer to chlorophyll in a functionally intact antenna protein. As can be seen from the comparison of the laser power density dependencies of ν4 band intensity and fluorescence intensity (Figure 7), the relatively steep decrease in the band intensity, observed at low laser power density values, is accompanied by a relatively small increase in the xanthophyll fluorescence quantum yield. On the other hand, a pronounced increase in the fluorescence yield of xanthophylls is observed at higher light intensities, at which the light-driven modulation of the ν4 band intensity is practically already at the saturation

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Figure 9. Fluorescence and RR spectrum of LHCII, recorded with laser power density 113 µW/µm2 (selected from Figure 6) presented along with components of Gaussian deconvolution (thick solid line), reconstituted spectrum (dashed line) representing the fluorescence spectrum, and residuals (thin solid line) representing the RR spectrum. Parameters of the Gaussian components: G1: center 16 273 cm-1 (full width at half-height 537 cm-1); G2: 17 007 cm-1 (788 cm-1); G3: 17 654 cm-1 (862 cm-1); G4: 18 277 cm-1 (778 cm-1); G5: 19 036 cm-1 (861 cm-1).

Figure 8. Comparison of the RR spectra of the xanthophylls (indicated) recorded in situ in LHCII, reported by Ruban et al.,24 with the difference spectrum calculated by subtraction of the LHCII spectra presented in Figure 5, corresponding to the laser power density of 11 and 113 µW/ µm2.

level. It is possible that the same group of pigments, which is involved in the light-driven molecular configuration changes, observed in the region of laser power density below 113 µW/ µm2, is also responsible for the slight fluorescence yield increase observed. On the other hand, it is rather unlikely that only this particular group of xanthophylls exclusively takes part in the pronounced increase in fluorescence emission, observed at high light power densities. In fact, the spectral characteristics of the light emission change (see Figure 6). A further increase in the xanthophyll fluorescence quantum yield is observed, at higher power density levels, at which the changes in intensity of the ν4 band are practically already saturated. This observation is consistent with the operation of light-driven molecular mechanisms that result in energetic uncoupling of accessory xanthophyll pigments and chlorophylls, reported in intact leaves4 and in isolated LHCII.3 The pronounced fluorescence yield increase may be also associated with strong light-induced damage of LHCII and unimpaired energy transfer from xanthophylls to chlorophylls.

The problem of LHCII stability and integrity during measurements is crucial for conclusions concerning physiological importance of light-induced configuration change of the proteinbound xanthophylls. Figure 10 presents laser power density dependency of intensity of the principal carotenoid RR band (ν1). A discrepancy from the linear relationship and lower than expected RR intensity observed at higher laser power density may indicate sample photobleaching. As can be seen, up to the power density of 113 µW/µm2 the dependency remains linear, which is an indication of photostability of the LHCII samples. Figure 11 presents electronic absorption spectra of LHCII before the experiments and after the experiments carried out with light power density of 57 and 1132 µW/µm2. Illumination of the sample for 10 s with 57 µW/µm2 results with a slight decrease in the light scattering signal (clearly observed in the longwavelength part of the spectrum because of the lack of absorption bands) and increased absorption in the Soret region of the spectrum. Such effects are indicative of decreased aggregation level of LHCII.27 Illumination of the sample with very strong light (1132 µW/µm2) is most probably associated with partial photodamage of the complex, which is indicated by the appearance of the short-wavelength absorbing spectral forms (below 400 nm) and relative decrease in the absorption in the spectral region typical of 0-0 vibronic transition of carotenoids (around 480 nm). The experiments analyzed in Figures 10 and 11 support the conclusion that at light power densities below 113 µW/µm2, at which light-driven change of

Light-harvesting Pigment-Protein Complex LHCII

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Figure 12. Resonance Raman spectra of LHCII presented in the ν4 band spectral region recorded with laser power density 3 µW/µm2 (marked “before”), afterward with 13 µW/µm2, and after 6 min dark adaptation again with 3 µW/µm2 (marked “after”). The spectra were normalized in the maximum of the ν1 band. The noise visible in the spectra recorded with very low laser power density has not been subjected to a smoothing procedure.

Figure 10. Dependence of intensity of the ν1 band of LHCII on power density of laser excitation beam applied. The dashed line presents the linear relationship characteristic of low power density values.

Figure 13. Resonance Raman spectra of LHCII presented in the ν4 band spectral region recorded with the laser power density 11 µW/ µm2. The samples contained different concentration of the detergent n-dodecyl-β-D-maltoside (DM), as indicated. The spectra were normalized in the maximum of the ν1 band.

Figure 11. Absorption spectra of LHCII before and after RR experiment performed with laser power density 57 and 1132 µW/µm2 (indicated). In order to record absorption spectra the samples were diluted with the buffer to yield absorbance value very close to 0.10 in the Qy band maximum (optical path 1 cm). Recorded absorption spectra were normalized in the Qy maximum in order to enable precise comparison.

configuration of the LHCII-bound xanthophyll(s) were observed, the pigments are stable and the complex is integral. Physiological importance of the light-driven molecular mechanism observed in LHCII in this (lower) range of light power densities

is further supported by the fact that light-dependent decrease in the ν4 band intensity, observed at 13 µW/µm2, is fully reversible after 6 min of dark adaptation (see Figure 12). A question that still remains open regards an exact physical mechanism directly responsible for the light-driven xanthophyll configuration changes in LHCII observed. One possible mechanism could be based on a transient, light-driven trans-cis isomerization of the xanthophyll. An observation relevant to support this concept can be the relatively intensive sub-band at 1135 cm-1, in the ν2 band region, visible in the spectra of LHCII but not in the spectra of the all-trans carotenoids (see Figures 3 and 4). This band is diagnostic of cis isomers of carotenoids.28 It is worth mentioning that a certain, small fraction of 13-cisand 9-cis-violaxanthin has been found in intact leaves29 and also in isolated LHCII6 and that the population of this fraction increased upon illumination. Another possible physical mech-

2512 J. Phys. Chem. B, Vol. 113, No. 8, 2009 anism responsible for the light-driven change of the xanthophyll configuration could be based on the thermo-optical effect proposed by Garab and co-workers to operate in LHCII.30,31 Although the laser power densities applied in our experiments are comparable with other resonance Raman studies reported, they are relatively high, and the effective number of light quanta absorbed by the sample at power density 11 µW/µm2 (calculated by integration of the 1 minus transmission spectrum) corresponds roughly to the photon flux density of 2000 µmol m-2 s-1 of sunlight. In our opinion, a relatively short time of exposure (10 s) makes our findings relevant from the physiological point of view and comparable with the effects in plants exposed to prolonged overexcitation. The observed light-induced change of molecular configuration of the LHCII-bound xanthophyll, tentatively assigned to violaxanthin, that results in weakening of the pigment-protein interaction (relaxed twisting) may have a sound physiological importance. Enzymatic de-epoxidation of violaxanthin to antheraxanthin and further on to zeaxanthin requires detachment of the pigment from the protein environment, referred to as making violaxanthin available for de-epoxidation, and has been postulated to be an important step of regulation of the activity of the xanthophyll cycle.32 It has been reported that acidification of the thylakoid interior alone is not sufficient to detach violaxanthin from the protein environment,18 but it is possible that the process of making violaxanthin available for enzymatic de-epoxidation is a process that is dependent on light-intensity, as originally proposed by Siefermann and Yamamoto.33 The light-driven changes of configuration of the LHCII-bound xanthophylls can possibly also influence the aggregation level of the complex11 (see also Figure 11), which has a pronounced effect on the photoprotection in the photosynthetic apparatus at the molecular level.13,34,35 Disaggregation of LHCII by the detergent treatment is known to modify the accessory pigment interaction to the protein environment and photosynthetic excitation energy transfer.27 Such a process is also associated with changes in the intensity of the resonance Raman scattering spectrum in the ν4 region of the xanthophylls that are in resonance at 514.5 nm (see Figure 13). On the other hand, the spectral changes observed in such a case are rather limited to the intensity decrease and not to a change in the shape of the band, as observed in the case of the light-driven effect discussed above. Acknowledgment. This research was financed by the Ministry of Science and Higher Education of Poland from the funds for science in the years 2008-2011 within the research project N N303 285034. The BIONAN network is also acknowledged for financial support. References and Notes (1) Liu, Z.; Yan, H.; Wang, K.; Kuang, T.; Zhang, J.; Gui, L.; An, X.; Chang, W. Nature 2004, 428, 287. (2) Ruban, A. V.; Lee, P. J.; Wentworth, M.; Young, A. J.; Horton, P. J. Biol. Chem. 1999, 274, 10458.

Gruszecki et al. (3) Gruszecki, W. I.; Kernen, P.; Krupa, Z.; Strasser, R. J. Biochim. Biophys. Acta 1994, 1188, 235. (4) Gruszecki, W. I.; Krupa, Z. Z. Naturforsch. 1993, 48c, 46. (5) Gruszecki, W. I.; Matula, M.; Naomi, K. C.; Koyama, Y.; Krupa, Z. Biochim. Biophys. Acta 1997, 1319, 267. (6) Grudzinski, W.; Matula, M.; Sielewiesiuk, J.; Kernen, P.; Krupa, Z.; Gruszecki, W. I. Biochim. Biophys. Acta 2001, 1503, 291. (7) Robert, B.; Horton, P.; Pascal, A. A.; Ruban, A. V. Trends Plant Sci. 2004, 9, 385. (8) Croce, R.; Cinque, G.; Holzwarth, A. R.; Bassi, R. Photosynth. Res. 2000, 64, 221. (9) Ruban, A. V.; Berera, R.; Ilioaia, C.; van Stokkum, I. H.; Kennis, J. T.; Pascal, A. A.; van Amerongen, H.; Robert, B.; Horton, P.; van Grondelle, R. Nature 2007, 450, 575. (10) Krupa, Z.; Huner, N.; Williams, J.; Maissan, E.; James, D. Plant Physiol. 1987, 84, 19. (11) Grudzinski, W.; Krupa, Z.; Garstka, M.; Maksymiec, W.; Swartz, T. E.; Gruszecki, W. I. Biochim. Biophys. Acta 2002, 1554, 108. (12) Gruszecki, W. I. Methods Mol. Biol. 2004, 274, 173. (13) Gruszecki, W. I.; Grudzinski, W.; Gospodarek, M.; Patyra, M.; Maksymiec, W. Biochim. Biophys. Acta 2006, 1757, 1504. (14) Gruszecki, W. I.; Grudzinski, W.; Matula, M.; Kernen, P.; Krupa, Z. Photosynth. Res. 1999, 59, 175. (15) Niedzwiedzki, D.; Krupa, Z.; Gruszecki, W. I. J. Photochem. Photobiol. B: Biol. 2005, 78, 109. (16) Allen, K. D.; Staehelin, L. A. Anal. Biochem. 1991, 194, 214. (17) Garstka, M.; Venema, J. H.; Rumak, I.; Gieczewska, K.; Rosiak, M.; Koziol-Lipinska, J.; Kierdaszuk, B.; Vredenberg, W. J.; Mostowska, A. Planta 2007, 226, 1165. (18) Ruban, A. V.; Pascal, A. A.; Robert, B.; Horton, P. J. Biol. Chem. 2001, 276, 24862. (19) Kupisz, K.; Sujak, A.; Patyra, M.; Trebacz, K.; Gruszecki, W. I. Biochim. Biophys. Acta 2008, 1778, 2334. (20) Milanowska, J.; Polit, A.; Wasylewski, Z.; Gruszecki, W. I. J. Photochem. Photobiol. B: Biol. 2003, 72, 1. (21) Niedz´wiedzki, D.; Gruszecki, W. I. Coll. Surf. B: Biointerfaces 2003, 28, 27.. (22) Rimai, L.; Heyde, M. E.; Gill, D. J. Am. Chem. Soc. 1973, 95, 4493. (23) Robert, B. The electronic structure, stereochemistry and resonance Raman spectroscopy of carotenoids In The Photochemistry of Carotenoids; Frank, H. A., Young, A. J., Britton, G., Cogdell, R. J., Eds.; Kluwer Academic Publishers.: Dordrecht, 1999; Vol. 8, pp 189. (24) Ruban, A. V.; Pascal, A.; Lee, P. J.; Robert, B.; Horton, P. J. Biol. Chem. 2002, 277, 42937. (25) Polivka, T.; Sundstrom, V. Chem. ReV. 2004, 104, 2021. (26) Josue, J. S.; Frank, H. A. J. Phys. Chem. A 2002, 106, 4815. (27) Voigt, B.; Krikunova, M.; Lokstein, H. Photosynth. Res. 2008, 95, 317. (28) Koyama, Y.; Fujii, R. Cis-Trans Carotenoids in Photosynthesis: Configurations, Excited-State Properties and Physiological Functions In The Photochemistry of Carotenoids; Frank, H. A., Young, A. J., Britton, G., Cogdell, R. J., Eds.; Kluwer Academic Publishers: Dordrecht, 1999; pp 161-188.. (29) Phillip, D.; Molnar, P.; Toth, G.; Young, A. J. J. Photochem. Photobiol. B: Biol. 1999, 49, 89. (30) Cseh, Z.; Vianelli, A.; Rajagopal, S.; Krumova, S.; Kovacs, L.; Papp, E.; Barzda, V.; Jennings, R.; Garab, G. Photosynth. Res. 2005, 86, 263. (31) Holm, J. K.; Varkonyi, Z.; Kovacs, L.; Posselt, D.; Garab, G. Photosynth. Res. 2005, 86, 275. (32) Hartel, H.; Lokstein, H.; Grimm, B.; Rank, B. Plant Physiol. 1996, 110, 471. (33) Siefermann, D.; Yamamoto, H. Y. Biochim. Biophys. Acta 1974, 357, 144. (34) Pascal, A. A.; Liu, Z.; Broess, K.; van Oort, B.; van Amerongen, H.; Wang, C.; Horton, P.; Robert, B.; Chang, W.; Ruban, A. Nature 2005, 436, 134. (35) Liu, C.; Zhang, Y.; Cao, D.; He, Y.; Kuang, T.; Yang, C. J. Biol. Chem. 2008, 283, 487.

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