Mechanism of QE-quenching


A.R. Crofts and Christine T. Yerkes

Program in Biophysics and Dept. of Microbiology, University of Illinois at Urbana-Champaign

We have studied energy-dependent fluorescence lowering (qE-quenching),and suggested a model to explain the experimental data currently available.The main elements of the model are: a) The qE-quenching reflects a mechanism associated with a component of the light-harvesting antenna rather than the reaction center of photosystem (PS) II; we suggest that it occurs through formation of an efficient quencher in one of the minor chlorophyll protein (CP) complexes. b) The minor CPs have glutamate residues instead of glutamines at positions shown in light harvesting complex II(LHCII) to be ligands to chlorophylls near the lumenal interface. We suggest that the quenching reflects a change in ligation of chlorophyll on protonation of these glutamate residues leading to formation of an exciton coupled dimer with a neighboring pigment, in which additional energy levels allow vibrational relaxation of the excited singlet. The model accounts for the dependence on low lumenal pH, the ligand residue changes between LHCII and the minor CPs, the preferential distribution of components of the xanthophyll cycle in the minor CPs, the inhibition of qE-quenching by DCCD, and the specific binding of DCCD to the minor CPs.

Fig. 1. Aligned sequences of minor CPs (CP29, CP26 and CP24) showing changes of glutamine chlorophyll-ligands to glutamates.

Fig.2. LHCII structure showing glutamine residues thought to be modified to glutamates in the minor CPs (CP29, CP26 and CP24).

Fluorescence lowering is a physiological device to dump exciton energy as heat before it reaches the reaction center of PS II. The mechanism must be poised so that quenching comes in before the lumenal pH falls low enough to inhibit the donor-side reactions. The mechanism involves a change in state of the antenna complex (with a pK in the range 5.5 in higher plant chloroplasts), leading to dissipation of the singlet state, and hence quenching of fluorescence. The formation of zeaxanthin is enhanced at low lumenal pH, but has a slower onset and longer decay, and thus represents a secondary process to cope with more extended exposure. Association of zeaxanthin (and possibly antheraxanthin) with CP29 (or the other minor LHCs), amplifies (or is required for) the effect of low lumenal pH in quenching the fluorescence. Structural models of the interface between PS II and the antenna, and fluorescence emission spectra, suggest that the minor complexes serve as "bridges" between the major LHCIIs and the reaction center antenna proteins. It thus seems likely, as suggested by Bassi and colleagues, that the main action in qE-quenching is at this interface, and involves CP29 (or the other minor LHCs), and not aggregation of LHCII as suggested by Horton and colleagues.

Molecular mechanism.

The mechanism of quenching could in principle be quite simple. Physico-chemical studies show that chlorophylls in solution at concentrations comparable to those in the leaf show no fluorescence, and this is attributed to an interaction between "statistical dimers" which introduces additional energy levels allowing thermal pathways for de-excitation. The chlorophylls in LHCII are held apart by ligation, with a variety of groups providing ligands. We assume a similar structure for CP24, CP26 and CP29 based on sequence homology.If an acidic chlorophyll ligand were accessible to H+ from the lumenal phase, the liganding properties would change on acidification. We suggest that such a change might allow the effected chlorophyll to interact at short enough range to form exciton-coupled bands with a neighboring chlorophyll or carotenoid, and thus form a quencher of fluorescence. A change in ligand properties could either lead to release of a chlorophyll previously bound, specific ligation of a previously loosely bound chlorophyll, or an exchange of ligands.

Sequence comparison and alignment can be used to identify the ligands in CP24, CP26 and CP29 by homology with the ligands identified in LHCII. Residue changes with appropriate properties are Q131E in CP29 (numbering as in Kuhlbrandt), and Q197E in CP24. Glutamines at both positions are fully conserved in LHCII. Other known ligands are conserved between the LHCII, CP29, CP26 and CP24 sequences, except for N183H in the Arabidopsis CP29, and in CP24s. Other residue changes which might affect ligation are the substitutions of P82V (Arabidopsis CP29) or P82G (CP24)(not shown).In the LHCII structure, the proline in helix B liberates the peptide >C=O group of G78 so that it can act as a ligand. None of these other ligand changes would be expected to lead to pK changes in the acid range. It should be noted that the Q131E change is also seen in LHCI sequences.

The structure for LHCII is shown in Fig. 2, with the location of the two glutamines and the chlorophylls they ligate labeled. Both residues are on the lumenal side of the structure, but in the hydrophobic domain; Q131 in particular is quite well buried. We assume similar locations for the glutamates identified as substitutes in the alignment above for CP24, CP26 or CP29. To account for the dependence of quenching on low lumenal pH, we would have to suggest that a channel exists to allow access of H+ to one or both of these glutamates from the lumen. In contrast to the glutamate ligands in LHCII, which form charge compensating pairs with arginine residues from elsewhere in the sequence (chlorophylls in italics in Fig. 1), there are no obvious conserved changes in the lumenal loops which might provide a similar compensating group for E197 (in CP24) or E131 (in CP29, CP26). The buried glutamates identified here might therefore be the sites at which DCCD reacts to block qE-quenching and might account for the preferential labeling by DCCD of the minor chlorophyll binding proteins, and also explain the effects of DCCD on protolytic processes associated with the donor-side reactions. In order for these residues to form stable adducts with DCCD, the H+-channel postulated above would have to allow access of H+, but not of H2O at high activity.This seems to be the case with the DCCD binding proteolipid (subunit cin E. coli) which is thought to contribute to a H+-channel through the F0-part of the ATP-synthase. If biochemical evidence confirms our suggestion, the LHCII molecular structure might provide important clues as to how a relatively bulky hydrophobic residue gains access to sites from which water is restricted.

Role of zeaxanthin

The conversion of violaxanthin to antheraxanthin and zeaxanthin in the minor CPs could enhance the quenching process either directly, by contributing to the quenching through triplet formation or through singlet mechanisms involving the energy level changes suggested by Owens et al., or indirectly through structural changes. In the Kühlbrandt structure, the luteins are in close proximity to both the chlorophylls ligated by the pertinent glutamines, or to potential dimer partners; if the components of the xanthophyll cycle occupy the same relative positions, interactions could readily occur, but it is premature to guess at what these might be.

Harry Frank and colleagues have recently made an extensive study of the energy levels in carotenoids, including those of the xanthophyll cycle. Carotenoids were previously believed to be non-fluorescent,- reports of fluorescence could be attributed to other contaminating pigments. With the advent of HPLC for purification, and sensitive fluorescence spectrometers,it has now been possible to measure fluorescence from ultra pure samples, and to determine quantum yields.
(From Frank, H., Chynwat, V., Desamero, R.Z.B., Farhoosh, R., Erickson,J. and Bautista, J. (1997) On the photophysics and photochemical properties of carotenoids and their role as light-harvesting pigments in photosynthesis.Pure and Applied Chemistry, in press. With thanks for copies of the Figs.below in GIF format.)

Energy level scheme of carotenoids. S0, S1and S2 are singlet states, T1 is the lowest lying triplet state.

Carotenoids have three main energy levels,- the ground state (S0),the S2 level, responsible for the absorption bands, and the S1. Fluorescence can occur from either the S2 or the S1 level, although the latter is rare in natural carotenoids.

The data for the S1 and S2 state energies of a series of spheroidene analog molecules having extents of p-electroncon jugation from 7 to 13 carbon-carbon double bonds, are fit by an A + B/(N + C) expression. The parameters were A = 3,802 cm-1, B = 1.1 x 105cm-1, and C = 0.500 for the series of S1-state energies, or C = 2.020 for the S2 state energies.

Relatively short synthetic carotenoids show fluorescence from the S1-level, and Frank and colleagues have used data from these, and extrapolation based on Hückel theory, and on energy gap law for radiationless transitions,to calculate energy levels and lifetimes.

A fit of the energy gap law to the S1 dynamics and energies of fucoxanthin (t), short synthetic carotenoids (s, u, v), mini-7-b-carotene(solid circles). From the curve (solid line) the energies of the S1-states of other carotenoids (open squares, including zeaxanthin (j) and violaxanthin (r)) were deduced.

From the perspective of green plant photosynthesis, and the energy transfer and protective roles of the carotenoids of the xanthophyll cycle, the interesting conclusions that come from these studies are as follows:

  1. The S1 levels of b-carotene (14,200cm-1) and zeaxanthin (14,188 cm-1) (with 10 double-bonds) are lower than that of chlorophyll a absorbing at 680 nm (14,705 cm-1). They cannot therefore act as light harvesting pigments, transferring energy to chlorophyll via the S1 state. They may, however, quench chlorophyll fluorescence by acting as acceptors of excitation.
  2. Epoxidation of zeaxanthin to form violaxanthin (9 double-bonds) raises the energy level of the S1 band (15,190 cm-1) to higher than chlorophyll a. Thus, in the "dark" state of the xanthophyll cycle, the violaxanthin can act as an energy donor, but conversion to zeaxanthin changes it to an acceptor, providing a rapid pathway for shunting the excitation energy from the chlorophylls to the environment as heat.

References

     
    1. Murata, N. and Sugahara, K. (1969) Biochim. Biophys. Acta, 189, 182-189.
    2. Wraight, C.A. and Crofts, A.R. (1970) Eur. J. Biochem. 17, 319-323.
    3. Krause, G.H. (1973) Biochim. Biophys. Acta 292, 715-728.
    4. Foyer, C., Furbank, R., Harbinson, J. and Horton, P. (1990) Photosynthe-sisResearch, 25, 83-100.
    5. Weis, E. and Berry, J. (1987) Biochim. Biophys. Acta, 894, 198-208.
    6. Genty, B., Briantais, J-M., and Baker, N.R. (1989) Biochim. Biophys. Acta990, 87-92.
    7. Krause, G.H. and Weis, E. (1991) Annu. Rev. Plant Physiol. Plant Mol. Biol.42, 313-349.
    8. Gilmore, A.M. and Yamamoto, H.Y. (1993) Proc. Natl. Acad. Sci. USA. 89,1899-1903.
    9. Hager, A. (1969) Planta 89, 224-243.
    10. Yamamoto, H.Y. and Kamite, L. (1972) Biochim. Biophys. Acta 267, 538-543.
    11. Demmig, B., and Björkman, O. (1987) Planta 171, 171-184.
    12. Ruban, A.V., Young, A.J., Pascal, A.A. and Horton, P. (1994) Plant Physiol.104, 227-234.
    13. Oxborough, K. and Horton, P. (1987) Photosynth. Res. 12, 119-128.
    14. Demmig-Adams, B. (1990) in Curr. Res. Photosyn., (Baltscheffsky, M., ed.),Vol. II, pp. 357-364, Kluwer Acad. Press, The Netherlands.
    15. Yamamoto, H.Y. (1979) Pure Appl. Chem. 51, 639-648.
    16. Crofts, A.R., Wraight, C.A. and Fleischman, D.(1971) FEBS Lett. 15, 89-100.
    17. Bowes, J.M. and Crofts, A.R. (1981) Biochim. Biophys. Acta, 637, 464-472.
    18. Schreiber, U. and Neubauer, C. (1990) Photosynth. Research 25, 279-293.
    19. Krieger, A., Moya, I. and  Weis, E. (1992) Biochim. Biophys. Acta 1102, 167-176.
    20. Vass, I.,  Styring, S., Hundal, T.,  Koivuniemi, A., Aro, E.M.and Andersson, B. (1992) Proc. Natl. Acad. Sci. USA. 89, 1408-1412.
    21. Nedbal, L.,  Samson, G. and Whitmarsh, J. (1992) Proc. Natl. Acad.Sci, USA. 89, 7929-7933.
    22. Meunier, P.C. and  Bendall, D.S. (1993) Photosynth. Research 37, 147-158.
    23. Rees, D., Young, A., Noctor, G., Britton, G. and Horton, P. (1989) FEBSLett. 256, 85-90.
    24. Horton, P., Ruban, A.V., Rees, D., Pascal, A.A., Noctor, G. and Young,A.J. (1991) FEBS Lett.  292, 1-4.
    25. Ruban, A.V. and Horton, P. (1992) Biochim. Biophys. Acta 1102, 30-38.
    26. Mullineaux, C.W., Pascal, A.A., Horton, P. and Holzwarth, A.R. (1993) Biochim.Biophys. Acta 1141, 23-28.
    27. Krieger, A., Moya, I. and Weis, E. (1992) Biochim. Biophys. Acta 1102,167-176
    28. Yerkes, C.T. and Crofts, A.R. (1992) In Research on Photosynthesis (Murata,N., ed.) Vol. II. pp. 635-638. Kluwer Academic Publ., Dorrdrecht.
    29. Dainese, P., Santini, C., Ghiretti-Magaldi, A. Marquardt, J., Tidu, V.,Mauro, S., Bergantino, E. and Bassi, R. (1992)  In Research on Photosynthesis(Murata, N., ed.) Vol. II. pp. 13-20. Kluwer Academic Publ., Dordrecht.
    30. Bratt, C.E. and Åkerlund, H.-E. (1992) In Research on Photosynthesis(Murata, N., ed.) Vol. I. pp. 231-234. Kluwer Academic Publ., Dordrecht.
    31. Green, B,R., Durnford, D. and Pichersky, E. (1992) In Research on Photosynthesis(Murata, N., ed.) Vol. I. pp. 195-202. Kluwer Academic Publ., Dordrecht.
    32. Owens, T.G., Shreve, A.P. and Albrecht, A.C. (1992) In Research on Photosynthesis(Murata, N., ed.) Vol. I. pp. 179-186. Kluwer Academic Publ., Dordrecht.
    33. Deamer, D. W., Crofts, A. R., and Packer, L. (1967).  Biochim. Biophys.Acta 131, 81-96.
    34. Krause, G.H. (1974) Biochim. Biophys. Acta 333, 301-313.
    35. Mullet, J.E. and Arntzen, C.J. (1980) Biochim. Biophys. Acta 589, 100-117
    36. Crofts, A. R., Deamer, D. W., and Packer, L. (1967)  Bio-phys. Acta,131, 97-118.
    37. Izawa, S., Connolly, T.N., Winget, G.D. and Good, N.E. (1967) Brookhaven.Symp. Biol. 19, 169-187.
    38. Drechsler, Z., Nelson, N. and Neumann, J. (1969) Biochim. Biophys. Acta189, 65-73.
    39. Karlish, S.J.D. and Avron, M. (1968) FEBS Lett. 1, 21-24.
    40. Jahns, P. and Junge, W. (1988) Eur. J. Biochem. 193, 731-736.
    41. Webber, A. and Gray, J.C. (1989) FEBS Lett. 249, 79-82.
    42. Ruban, A.V., Walters, R.G. and Horton, P. (1992) FEBS Lett. 309, 175-179.
    43. Kühlbrandt, W., Wang, D.N. and Fujiyoshi, Y. (1984) Nature (Lond.)367, 614-621.
    44. Bassi, R., Pineau, B., Dianese, P. and Marquardt, J. (1993) European J.Biochem. 212, 297-303.
    45. Gilmore, A.M. and Yamamoto, H.Y. (1993) Photosynth. Research.  35,67-78.
    46. Chen, G.X.,  Kazimir, J. and Cheniae, G.M. (1992) Biochemistry, 31,11072-11083.
    47. Yerkes,  C.T. and Crofts, A.R. (1992) EBEC Short Reports, 7, 5.
    48. Beddard, G.S., Carlin, S.E. and Porter, G. (1976) Chem. Phys. Lett. 43,27-32.
    49. Jennings, R.C., Zucchelli, G., Bassi, R., Vianelli, A. and Garlaschi, F.(1994) Biochim. Biophys. Acta 1184, 279-283.