Title: Configuration and Dynamics of Xanthophylls in Light-harvesting Antennae of Higher Plants
Abstract: Resonance Raman excitation spectroscopy combined with ultra low temperature absorption spectral analysis of the major xanthophylls of higher plants in isolated antenna and intact thylakoid membranes was used to identify carotenoid absorption regions and study their molecular configuration. The major electronic transitions of the light-harvesting complex of photosystem II (LHCIIb) xanthophylls have been identified for both the monomeric and trimeric states of the complex. One long wavelength state of lutein with a 0-0 transition at 510 nm was detected in LHCIIb trimers. The short wavelength 0-0 transitions of lutein and neoxanthin were located at 495 and 486 nm, respectively. In monomeric LHCIIb, both luteins absorb around 495 nm, but slight differences in their protein environments give rise to a broadening of this band. The resonance Raman spectra of violaxanthin and zeaxanthin in intact thylakoid membranes was determined. The broad 0-0 absorption transition for zeaxanthin was found to be located in the 503–511 nm region. Violaxanthin exhibited heterogeneity, having two populations with one absorbing at 497 nm (0-0), 460 nm (0-1), and 429 nm (0-2), and the other major pool absorbing at 488 nm (0-0), 452 nm (0-1), and 423 nm (0-2). The origin of this heterogeneity is discussed. The configuration of zeaxanthin and violaxanthin in thylakoid membranes was different from that of free pigments, and both xanthophylls (notably, zeaxanthin) were found to be well coordinated within the antenna proteins in vivo, arguing against the possibility of their free diffusion in the membrane and supporting our recent biochemical evidence of their association with intact oligomeric light-harvesting complexes (Ruban, A. V., Lee, P. J., Wentworth, M., Young, A. J., and Horton, P. (1999) J. Biol. Chem. 274, 10458–10465). Resonance Raman excitation spectroscopy combined with ultra low temperature absorption spectral analysis of the major xanthophylls of higher plants in isolated antenna and intact thylakoid membranes was used to identify carotenoid absorption regions and study their molecular configuration. The major electronic transitions of the light-harvesting complex of photosystem II (LHCIIb) xanthophylls have been identified for both the monomeric and trimeric states of the complex. One long wavelength state of lutein with a 0-0 transition at 510 nm was detected in LHCIIb trimers. The short wavelength 0-0 transitions of lutein and neoxanthin were located at 495 and 486 nm, respectively. In monomeric LHCIIb, both luteins absorb around 495 nm, but slight differences in their protein environments give rise to a broadening of this band. The resonance Raman spectra of violaxanthin and zeaxanthin in intact thylakoid membranes was determined. The broad 0-0 absorption transition for zeaxanthin was found to be located in the 503–511 nm region. Violaxanthin exhibited heterogeneity, having two populations with one absorbing at 497 nm (0-0), 460 nm (0-1), and 429 nm (0-2), and the other major pool absorbing at 488 nm (0-0), 452 nm (0-1), and 423 nm (0-2). The origin of this heterogeneity is discussed. The configuration of zeaxanthin and violaxanthin in thylakoid membranes was different from that of free pigments, and both xanthophylls (notably, zeaxanthin) were found to be well coordinated within the antenna proteins in vivo, arguing against the possibility of their free diffusion in the membrane and supporting our recent biochemical evidence of their association with intact oligomeric light-harvesting complexes (Ruban, A. V., Lee, P. J., Wentworth, M., Young, A. J., and Horton, P. (1999) J. Biol. Chem. 274, 10458–10465). light-harvesting antenna resonance Raman photosystem II light-harvesting complex difference spectrum for zeaxanthin and violaxanthin The light-harvesting antenna (LHA)1 of higher plants binds five types of xanthophylls: lutein, neoxanthin, violaxanthin, zeaxanthin, and antheraxanthin. The last three constitute the xanthophyll cycle, which has been suggested to participate in the process of dissipation of excess excitation energy, giving rise to nonphotochemical fluorescence quenching (1Demmig-Adams B. Biochim. Biophys. Acta. 1990; 1020: 1-24Crossref Scopus (1377) Google Scholar, 2Demmig-Adams B. Adams III, W.W. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1992; 43: 599-626Crossref Scopus (2045) Google Scholar, 3Horton P. Ruban A.V. Walters R.G. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1996; 47: 65-84Crossref Scopus (1398) Google Scholar). Several important questions concerning these xanthophylls remain unanswered: why is there such a variety of xanthophyll types in antenna; what is the exact molecular mechanism of zeaxanthin action in nonphotochemical fluorescence quenching; where are xanthophylls located; and what is the nature of their interaction with the protein and other pigments? LHA consists of a number of pigment-protein complexes accommodating different types and amounts of xanthophylls (5Peter G.F. Thornber P. J. Biol. Chem. 1991; 266: 16745-16754Abstract Full Text PDF PubMed Google Scholar, 6Bassi R. Pineau B. Dainese P. Marquardt J. Eur. J. Biochem. 1993; 212: 297-303Crossref PubMed Scopus (349) Google Scholar, 7Sandora D. Croce R. Pagano A. Crimi M. Bassi R. Biochim. Biophys. Acta. 1998; 1365: 207-214Crossref PubMed Scopus (95) Google Scholar, 8Ruban A.V. Young A.J. Pascal A.A. Horton P. Plant Physiol. 1994; 104: 227-234Crossref PubMed Scopus (212) Google Scholar). The major and most characterized LHA complex, the trimeric LHCIIb, binds 2 luteins, 1 neoxanthin, and between 0.1 and 1 violaxanthin per monomer (7Sandora D. Croce R. Pagano A. Crimi M. Bassi R. Biochim. Biophys. Acta. 1998; 1365: 207-214Crossref PubMed Scopus (95) Google Scholar, 9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). The amount of bound violaxanthin was found to depend on the treatment during purification (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar), as well as on plant growth conditions (10Verhoeven A.S. Adams W.W. Demmig-Adams B. Croce R. Bassi R. Plant Physiology. 1999; 120: 727-737Crossref PubMed Scopus (98) Google Scholar). The two luteins of LHCIIb are thought to correspond to the two carotenoid molecules located near the transmembrane helixes A and B in the inner core of the complex (11Kuhlbrandt W. Wang D.N. Fujiyoshi Y. Nature. 1994; 367: 130-135Crossref Scopus (1529) Google Scholar), thus being tightly associated with it and probably having a structural role. Site-directed mutagenesis experiments have suggested that neoxanthin is associated with helix C (12Croce R. Remelli R. Varotto C. Breton J. Bassi R. FEBS Lett. 1999; 456: 1-6Crossref PubMed Scopus (104) Google Scholar). Neoxanthin was found to have the highest affinity of binding to the complex (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). In contrast, the binding affinity of violaxanthin to the complex is the lowest, and it can be easily removed by various treatments. This biochemical work has therefore established that a large population of xanthophyll cycle carotenoids is peripherally bound to LHA complexes.The structure of the minor PSII antenna complexes as well as all LHCI complexes is not known; therefore, it is not clear where xanthophyll molecules are bound to them, what the nature of this binding is, or what the carotenoid configuration is involved. The data on xanthophyll stoichiometry in the minor PSII antenna are controversial (4Lee A.L. Thornber J.P. Plant Physiol. 1995; 85: 495-502Google Scholar, 6Bassi R. Pineau B. Dainese P. Marquardt J. Eur. J. Biochem. 1993; 212: 297-303Crossref PubMed Scopus (349) Google Scholar, 8Ruban A.V. Young A.J. Pascal A.A. Horton P. Plant Physiol. 1994; 104: 227-234Crossref PubMed Scopus (212) Google Scholar) and may well again reflect natural variation of xanthophyll ratios. However, it is clear that unlike LHCIIb, the minor antenna PSII complexes, CP24, CP26, and particularly CP29, contain at least one strongly bound xanthophyll cycle carotenoid. The efficiency of violaxanthin de-epoxidation—a prerequisite for the photoprotective energy dissipation state in thylakoid—was found to be the reciprocal of the relative affinity of binding of this xanthophyll to various antenna components, suggesting that only loosely bound molecules are accessible to the violaxanthin de-epoxidase (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). A similar tendency was observed for the PSI antenna, which contains at least 50% tightly bound violaxanthin that was not converted into zeaxanthin even under conditions favoring maximum de-epoxidation (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar, 13Farber A. Young A.J. Ruban A.V. Horton P. Jahns P. Plant Physiol. 1997; 115: 1609-1618Crossref PubMed Scopus (173) Google Scholar). Therefore, it may be argued that the "structural" violaxanthin molecules, which are strongly coordinated within the minor LHA complexes, do not play a role in the xanthophyll cycle.It is clear that new methodologies are needed to identify, locate, and analyze xanthophylls and their configuration and function in LHA complexes. One powerful approach has been to reconstitute light-harvesting complexes from Lhcb polypeptides and pigment mixtures of varying composition. Site-directed mutagenesis of these polypeptides has been used to determine the location and specificity of carotenoid binding sites (12Croce R. Remelli R. Varotto C. Breton J. Bassi R. FEBS Lett. 1999; 456: 1-6Crossref PubMed Scopus (104) Google Scholar). The effects of carotenoid binding to peripheral sites in LHA complexes has allowed investigation of their role in energy dissipation (14Ruban A.V. Young A. Horton P. Biochim. Biophys. Acta-Bioenerg. 1994; 1186: 123-127Crossref Scopus (112) Google Scholar, 15Ruban A.V. Young A.J. Horton P. Biochemistry. 1996; 35: 674-678Crossref PubMed Scopus (115) Google Scholar, 16Ruban A.V. Phillip D. Young A.J. Horton P. Biochemistry. 1997; 36: 7855-7859Crossref PubMed Scopus (110) Google Scholar, 17Wentworth M. Ruban A.V. Horton P. FEBS Lett. 2000; 471: 71-74Crossref PubMed Scopus (62) Google Scholar). Along with the development of these biochemical techniques there has been progress toward the development of instrumental methods to analyze antenna xanthophylls. Absorption, linear and circular dichroism, and triplet state spectroscopies have been applied to identify their electronic absorption bands and energy transfer pathways to and from chlorophyll molecules (18Peterman E.J.G. Gradinaru C.C. Calkoen F. Borst J.C. van Grondelle R. van Amerongen H. Biochemistry. 1997; 36: 12208-12215Crossref PubMed Scopus (102) Google Scholar, 19Peterman E.J.G. Monshouwer R. van Stokkum I.H.M. van Grondelle R. van Amerongen H. Chem. Phys. Lett. 1997; 264: 279-284Crossref Scopus (53) Google Scholar, 20Ruban A.V. Calkoen F. Kwa S.L.S. van Grondelle R. Horton P. Dekker J.P. Biochim. Biophys. Acta. 1997; 1321: 61-70Crossref Scopus (105) Google Scholar, 21Connelly J.P. Muller M.G. Hucke M. Gatzen G. Mullineaux C.W. Ruban A.V. Horton P. Holzwarth A.R. J. Phys. Chem. B. 1997; 101: 1902-1909Crossref Scopus (107) Google Scholar, 22Gradinaru C.C. van Stokkum I.H.M. Pascal A.A. van Grondelle R. van Amerongen H. J. Phys. Chem. B. 2000; 104: 9330-9342Crossref Scopus (181) Google Scholar). However, it remains difficult to make unambiguous assignments for even simpler systems, such as those containing only three xanthophylls, and impossible to make reasonable spectral assignments in the Soret absorption region of whole thylakoid membranes. Recently, we have combined optical absorption spectroscopy with selective excitation resonance Raman spectroscopy in order to identify the absorption transitions of lutein and neoxanthin in LHCIIb trimers (23Ruban A.V. Pascal A.A. Robert B. FEBS Lett. 2000; 477: 181-185Crossref PubMed Scopus (97) Google Scholar). Unlike previous work (18Peterman E.J.G. Gradinaru C.C. Calkoen F. Borst J.C. van Grondelle R. van Amerongen H. Biochemistry. 1997; 36: 12208-12215Crossref PubMed Scopus (102) Google Scholar), which used LHCIIb containing three types of carotenoids, we have prepared LHCII, which contained only lutein and neoxanthin, a more simple system. The characteristic Raman feature for neoxanthin and lutein, the ν1 maximum position, was identified and used to build the Raman excitation profiles for the isolated complex. This allowed not only an estimation of the energies for the three absorption bands (0-0, 0-1, and 0-2) but also an observation of the dynamics of carotenoid configuration upon oligomerization of the complex.The approaches mentioned above are susceptible to various artifacts and limitations, such as the removal of lipids and pigments by the detergents used in the isolation and reconstitution procedures and the resulting alteration in protein conformation, xanthophyll binding affinity, and xanthophyll environment. One way forward would be the isolation of a more integrated LHA, in which pigments are less perturbed by detergent treatment. For example, we have recently isolated an oligomeric LHCIIb-enriched antenna complex using very mild detergent treatment of PSII particles and unstacked thylakoid membranes (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar), and this preparation contained a higher proportion of violaxanthin and zeaxanthin. However, the most ideal approach would be to analyze carotenoid structure and dynamics in situ in intact thylakoid membranes.The aim of the work described in this paper is to further develop the new spectroscopic methodologies to analyze xanthophylls, and most importantly to make progress toward their application to more complex systems including the whole thylakoid membrane. This approach not only enables the determination of the electronic transitions of all xanthophylls in vivo but also allows monitoring of their conformational and configurational dynamics and binding within the native LHA. To achieve this aim, we have first undertaken a systematic search for the characteristic features of isolated carotenoids that are found in the photosynthetic membrane, in order to use them for constructing the resonance Raman profiles. This allowed absorption band assignments not only for xanthophylls of isolated antenna complexes but also violaxanthin and zeaxanthin of intact antennae in thylakoid membranes. We have obtained for the first time the resonance Raman spectra of violaxanthin and zeaxanthin in vivo. It is concluded that zeaxanthin adopts a configuration that is likely to reflect its well defined binding within the antenna, rather than a free location in the membrane. This work establishes a new approach to the study of complex carotenoid-containing systems and offers a broad range of applications, from identification and assessment of xanthophyll configuration in reconstituted/isolated complexes to in vivoinvestigation of xanthophyll cycle carotenoids in order to establish their role in photoprotective mechanisms.MATERIALS AND METHODSCarotenoid samples were prepared as described in Ref. 24Phillip D. Ruban A.V. Horton P. Asato A. Young A.J. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 1492-1497Crossref PubMed Scopus (63) Google Scholar by Dr. Denise Phillip (John Moores University, Liverpool, United Kingdom). LHCIIb was prepared from dark-adapted spinach leaves using isoelecrofocusing of PSII-enriched particles, as described in Ref. 8Ruban A.V. Young A.J. Pascal A.A. Horton P. Plant Physiol. 1994; 104: 227-234Crossref PubMed Scopus (212) Google Scholar. Purification of LHCIIb trimers and removal of violaxanthin was carried out on a sucrose gradient (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). LHCIIb monomers were prepared by phospholipase A2 treatment of LHCIIb trimers for 36 h in the presence of 20 mm of CaCl2 at a chlorophyll concentration of 0.5 mm, followed by purification on a sucrose gradient as described previously (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). Intact thylakoid membranes were obtained by the procedure described in Ref. 8Ruban A.V. Young A.J. Pascal A.A. Horton P. Plant Physiol. 1994; 104: 227-234Crossref PubMed Scopus (212) Google Scholar. To induce maximum violaxanthin de-epoxidation, thylakoids were incubated at room temperature at a chlorophyll concentration of 200 μm for 2 h in a medium containing 5 mmd-isoascorbate, 10 mm HEPES, and 10 mm sodium citrate at pH 5.5 with or without 5 mm Mg2+.Absorption spectra were recorded on a Varian Cary E5 double-beam scanning spectrophotometer; measurements at 4 K were performed using a helium bath cryostat (Utreks). Low temperature resonance Raman spectra were obtained in a helium flow cryostat (Air Liquide, Paris, France) using a Jobin-Yvon U1000 Raman spectrophotometer equipped with a liquid nitrogen-cooled charge-coupled devices detector (Spectrum One, Jobin-Yvon, Paris, France) as described in Ref.25Sturgis J.N. Robert B. J. Mol. Biol. 1994; 238: 445-454Crossref PubMed Scopus (78) Google Scholar. Excitation was provided by Coherent Argon (Innova 100) and Krypton (Innova 90) lasers (at 457.9, 476.5, 496.5, 488.0, 501.7, and 514.5 nm and at 528.7 and 413.1 nm, respectively) and a Liconix helium-cadmium laser (at 441.6 nm). The choice of this wavelength range was determined by the absorption profiles of the xanthophylls used. Fig. 1 displays absorption spectra of the four major xanthophylls that have been studied. The number of laser excitation lines (indicated in Fig. 1 by dotted arrows) covers all of the 0-0 transitions and also part of the electron-vibrational bands (0-1 and 0-2). It is likely that carotenoid absorption maxima in vivo will be somewhat shifted from those in organic solvent. However, from previous experience with LHCIIb xanthophylls, we have found that pyridine provides an environment with polarizability (n = 1.5092) much closer to that of lipid and membrane protein environments than such commonly used solvents as n-hexane (n = 1.3750) or ethanol (1.3611). The amplitude of the resonance effect is proportional to the square of the absorption probability. This offers an efficient line-narrowing effect in the Raman excitation profiles compared with absorption spectra, and the increased selectivity makes it possible to resolve complex bandshapes.DISCUSSIONIn this paper, we have demonstrated a new approach to the characterization of higher plant xanthophylls using comparison of absorption band structure and resonance Raman excitation profiles. The approach proved to be an effective methodology for identification and monitoring of the molecular conformation and configuration of carotenoids both in isolated pigment-protein complexes and, most significantly, in intact thylakoid membranes. This method is based on the identification of a number of characteristic fingerprints in the resonance Raman spectrum for each carotenoid involved. Analysis of isolated LHCIIb monomers and trimers containing only two types of xanthophylls, lutein and neoxanthin, allowed the identification of their corresponding absorption bands and molecular configuration.In the monomeric state of LHCIIb, the configuration and environment of the two LHCIIb luteins are very similar, both absorbing at 495 nm. The broader full width at half maximum for the band at 495 nm in the monomer spectrum compared with that of the trimer (Fig. 4) may suggest that their maxima positions differ within this region and therefore that their environment is also slightly different. In the trimer there is a 510 nm band not found in the monomer; this band was assigned to a red-shifted lutein in the former. Its intensity was significantly reduced in the second derivative spectrum because of the much broader FWHM compared with the short wavelength form of the pigment. In the trimer, the 510 nm lutein undergoes significant twisting, as a result either of an influence of the protein or of interaction with the other pigments or lipids present. It is not clear from our data whether this lutein plays a specific role in, for example, stabilization of the trimer and/or photoprotection of chlorophylls. However, these large differences in environment of the two luteins are important from structural and spectroscopic viewpoints (11Kuhlbrandt W. Wang D.N. Fujiyoshi Y. Nature. 1994; 367: 130-135Crossref Scopus (1529) Google Scholar, 18Peterman E.J.G. Gradinaru C.C. Calkoen F. Borst J.C. van Grondelle R. van Amerongen H. Biochemistry. 1997; 36: 12208-12215Crossref PubMed Scopus (102) Google Scholar, 19Peterman E.J.G. Monshouwer R. van Stokkum I.H.M. van Grondelle R. van Amerongen H. Chem. Phys. Lett. 1997; 264: 279-284Crossref Scopus (53) Google Scholar, 20Ruban A.V. Calkoen F. Kwa S.L.S. van Grondelle R. Horton P. Dekker J.P. Biochim. Biophys. Acta. 1997; 1321: 61-70Crossref Scopus (105) Google Scholar, 21Connelly J.P. Muller M.G. Hucke M. Gatzen G. Mullineaux C.W. Ruban A.V. Horton P. Holzwarth A.R. J. Phys. Chem. B. 1997; 101: 1902-1909Crossref Scopus (107) Google Scholar). This approach has revealed the molecular dynamics of a bound xanthophyll as a function of the state of oligomerization of the complex, and this for the first time. These factors may (at least in part) be behind the existence of monomeric (minor antenna), trimeric (major LHCIIb), and oligomeric states of antenna proteins in vivo. In addition, the methodology described here could be also used in the investigation of the LHC reconstitution process, using various types of xanthophylls (33Giuffra E. Cugini D. Croce R. Bassi R. Eur. J. Biochem. 1996; 238: 112-120Crossref PubMed Scopus (116) Google Scholar, 34Paulsen H. Frank H.A. Young A.J. Britton G. Cogdell R.J. The Photochemistry of Carotenoids. Kluwer Academic Publishers, The Netherlands1999: 123-135Google Scholar).The application of RR spectroscopy to thylakoid membranes has revealed important new information about the xanthophyll cycle carotenoids, zeaxanthin and violaxanthin. The physiological role of these carotenoids is unclear, as is their location in the thylakoid membrane. Indirect measurements (correlating nonphotochemical fluorescence quenching with de-epoxidation state) have suggested that xanthophyll cycle carotenoids may be completely free in the thylakoid membrane and can only move to a very small number of quenching centers in LHCII proteins upon ΔpH formation (35Gilmore A. Physiol. Plant. 1997; 99: 197-205Crossref Google Scholar). Some data on isolated LHCII have suggested that only the minor LHCII can bind violaxathin and zeaxanthin tightly and that therefore these must be the functional proteins in nonphotochemical fluorescence quenching (36Bassi R. Croce R. Cugini D. Sandona D. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 10056-10061Crossref PubMed Scopus (189) Google Scholar). In contrast, we have suggested that all of these carotenoids are bound, mostly to peripheral sites on each of the proteins. The strongly associated violaxanthin molecules located within minor complexes were found to be inaccessible to the de-epoxidase enzyme and therefore unlikely to be involved in photoprotection (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar, 13Farber A. Young A.J. Ruban A.V. Horton P. Jahns P. Plant Physiol. 1997; 115: 1609-1618Crossref PubMed Scopus (173) Google Scholar). Therefore, we have suggested that the peripheral violaxanthin and zeaxanthin molecules play the key role in photoprotective energy dissipation. In this study, we have been able to obtain the first data on the state of the xanthophyll cycle carotenoids in the thylakoid membrane. The more structured RR spectrum for both violaxanthin and zeaxanthin in the thylakoid membrane compared with those free in solution indicate that both of these carotenoids are in fact in well coordinated environments in the thylakoid membrane, almost certainly bound to protein and not free to move. Zeaxanthin appeared to be more distorted than violaxanthin. Indeed, the ν4region is more structured and intense for zeaxanthin than violaxanthin, suggesting that the former is in a tight association with the protein, consistent with estimations of binding affinities to LHCII proteins (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar).Ultra-low temperature absoprtion spectroscopy has also revealed new information about these carotenoids. The zeaxanthin-minus-violaxanthin absorption spectrum of thylakoids had a complex structure. The heterogeneity within this spectrum suggested that violaxanthin exists in two different populations. This suggestion is consistent with the observation that violaxanthin was found to be differently bound to the minor and major LHCII and LHCI complexes. It is possible that one population corresponds to violaxanthin associated with the minor LHCII or LHCI, whereas the other is peripherally bound to LHCIIb. An alternative explanation is that the dual band structure of the difference spectrum zeaxanthin minus violaxanthin originates from excitonically coupled violaxanthin. In this case, the coupling energy (V) will be within 180 cm−1 taking into account the Davidov's splitting (E) of approximately 360 cm−1 (E = 2·V). In the case of a moderate transition dipole moment and a parallel orientation of two interacting violaxanthin molecules, they should be located within 0.5–0.7 nm of each other (37Davidov A.S. Theory of Molecular Excitons. McGraw-Hill, New York1962Google Scholar, 38Kenkre V.M. Knox R.S. Phys. Rev. Lett. 1974; 33: 803-806Crossref Scopus (127) Google Scholar).It is interesting to mention that acidification did not cause alteration in violaxanthin maximum positions, which one would anticipate in the case of ΔpH-induced detachment of violaxanthin from antenna to undergo de-epoxidation. Resonance Raman spectra were also found to be unaffected (data not shown). This indicates that acidification has no immediate effect on the state of violaxanthin.Clearly, the application of spectroscopic methods to the analysis of xanthophylls in isolated complexes and intact thylakoids provides a new approach to understanding the structure and dynamics of these molecules. In particular, the selectivity of the RR technique and its freedom from the artifacts and problems normally associated with absorption and fluorescence measurements on complex intact systems have provided new information about violaxanthin and zeaxanthin in vivo. In the future, we hope to apply a similar methodology to whole leaves.AcknowlegdementsWe acknowledge Dr. Denise Phillip and Prof. Andrew Young for a kind gift of carotenoid samples. The light-harvesting antenna (LHA)1 of higher plants binds five types of xanthophylls: lutein, neoxanthin, violaxanthin, zeaxanthin, and antheraxanthin. The last three constitute the xanthophyll cycle, which has been suggested to participate in the process of dissipation of excess excitation energy, giving rise to nonphotochemical fluorescence quenching (1Demmig-Adams B. Biochim. Biophys. Acta. 1990; 1020: 1-24Crossref Scopus (1377) Google Scholar, 2Demmig-Adams B. Adams III, W.W. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1992; 43: 599-626Crossref Scopus (2045) Google Scholar, 3Horton P. Ruban A.V. Walters R.G. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1996; 47: 65-84Crossref Scopus (1398) Google Scholar). Several important questions concerning these xanthophylls remain unanswered: why is there such a variety of xanthophyll types in antenna; what is the exact molecular mechanism of zeaxanthin action in nonphotochemical fluorescence quenching; where are xanthophylls located; and what is the nature of their interaction with the protein and other pigments? LHA consists of a number of pigment-protein complexes accommodating different types and amounts of xanthophylls (5Peter G.F. Thornber P. J. Biol. Chem. 1991; 266: 16745-16754Abstract Full Text PDF PubMed Google Scholar, 6Bassi R. Pineau B. Dainese P. Marquardt J. Eur. J. Biochem. 1993; 212: 297-303Crossref PubMed Scopus (349) Google Scholar, 7Sandora D. Croce R. Pagano A. Crimi M. Bassi R. Biochim. Biophys. Acta. 1998; 1365: 207-214Crossref PubMed Scopus (95) Google Scholar, 8Ruban A.V. Young A.J. Pascal A.A. Horton P. Plant Physiol. 1994; 104: 227-234Crossref PubMed Scopus (212) Google Scholar). The major and most characterized LHA complex, the trimeric LHCIIb, binds 2 luteins, 1 neoxanthin, and between 0.1 and 1 violaxanthin per monomer (7Sandora D. Croce R. Pagano A. Crimi M. Bassi R. Biochim. Biophys. Acta. 1998; 1365: 207-214Crossref PubMed Scopus (95) Google Scholar, 9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar). The amount of bound violaxanthin was found to depend on the treatment during purification (9Ruban A.V. Lee P.J. Wentworth M. Young A.J. Horton P. J. Biol. Chem. 1999; 274: 10458-10465Abstract Full Text Full Text PDF PubMed Scopus (222) Google Scholar), as well as on plant growth condit