From the Institut für Biochemie der Pflanzen,
Heinrich-Heine-Universität Düsseldorf, D-40225
Düsseldorf, Germany and ¶ Institut für Allgemeine
Botanik, Johannes-Gutenberg-Universität Mainz, Müllerweg 6, D-55128 Mainz, Germany
Received for publication, March 9, 2001, and in revised form, March 30, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In higher plants, the de-epoxidation of
violaxanthin (Vx) to antheraxanthin and zeaxanthin is required for the
pH-dependent dissipation of excess light energy as heat and
by that process plays an important role in the protection
against photo-oxidative damage. The de-epoxidation reaction was
investigated in an in vitro system using reconstituted
light-harvesting complex II (LHCII) and a thylakoid raw extract
enriched in the enzyme Vx de-epoxidase. Reconstitution of LHCII with
varying carotenoids was performed to replace lutein and/or neoxanthin,
which are bound to the native complex, by Vx. Recombinant LHCII
containing either 2 lutein and 1 Vx or 1.6 Vx and 1.1 neoxanthin or 2.8 Vx per monomer were studied. Vx de-epoxidation was inducible for all
complexes after the addition of Vx de-epoxidase but to different
extents and with different kinetics in each complex. Analysis of the
kinetics indicated that the three possible Vx binding sites have at
least two, and perhaps three, specific rate constants for
de-epoxidation. In particular, Vx bound to one of the two lutein
binding sites of the native complex, most likely L1, was not at all or
only at a slow rate convertible to Zx. In reisolated LHCII, newly
formed Zx almost stoichiometrically replaced the transformed Vx,
indicating that LHCII and Vx de-epoxidase stayed in close
contact during the de-epoxidation reactions and that no release of
carotenoids occurred.
Antenna proteins serve as light-harvesting systems in all
photosynthetic organisms that collect energy for the primary light reactions in the reaction centers. In higher plants, the antenna proteins of both photosystems constitute the large family of
light-harvesting chlorophyll
(Chl)1 a/b-binding (LHC)
proteins (1). All LHC proteins are encoded by nuclear Lhc
genes, show high sequence similarity among themselves (1-3), and have
presumably similar structures (1). The structure of the major LHC of
photosystem II (PS II), the so-called LHCII or Lhcb1/2 protein, has
been determined at 3.4 Å by electron crystallography (5). According to
biochemical analysis, LHCII binds two lutein (Lu), 1 neoxanthin (Nx),
and substoichiometric amounts of violaxanthin (Vx)/monomer (6-8). Two
xanthophylls, which are associated with the two central
membrane-spanning The selectivity of the three carotenoid binding sites in LHCII has been
studied using recombinant protein (11, 12). It was shown that Vx could
bind to all binding sites of the native complex, the two Lu (L1 and L2)
as well as the Nx binding site, albeit with lower affinity than Lu and
Nx. In the absence of other carotenoids during reconstitution, 2-3 Vx
were found to bind to LHCII (11, 12). In the presence of equal amounts
of Lu and Vx, roughly 2 Lu and 1 Vx bound to the complexes (11, 12).
The dissipation of excess light energy in the antenna of PSII,
frequently measured as non-photochemical quenching (NPQ) of Chl
fluorescence, is supposed to be important for the protection of plants
against photo-oxidative damage (13-15). The de-epoxidation of Vx to
zeaxanthin (Zx) via the intermediate antheraxanthin (Ax) in the
xanthophyll cycle plays a crucial role in the
Preparation of Pigments, Protein, and
Reconstitution--
Pigments and overexpressed Lhcb1*2 were isolated
as described (12). Reconstitution was performed by detergent exchange
as described (23) at a Chl a/b ratio of 3 and a Chl and xanthophyll concentration of 0.8 and 0.1 mg/ml, respectively. Xanthophylls were
added either as single species or as 1:1 mixtures. Vx/Zx ratios for
reconstitution of Nx·Vx·Zx complexes were as depicted in
Fig. 2. Because reconstituted LHCII was separated from unbound pigments
by sucrose density centrifugation, sucrose was omitted from the
solubilization buffer. 200 µl of reconstitution sample were applied
to 11 ml of sucrose gradients, generated by one freeze-thaw cycle (0.4 M sucrose, 5 mM Tricine-NaOH, pH 7.8, 0.05%
(w/v) dodecylmaltoside), and centrifuged at 280,000 × g for 17 h at 4 °C.
Pigment Analysis--
Pigments were extracted from all samples
following the method of (24) and separated by reversed-phase HPLC as
described in (18).
Preparation of Crude VxDE Extracts--
VxDE extracts were
isolated from spinach essentially following the procedure described by
Arvidsson et al. (25). Roughly, isolated thylakoids were
broken by sonification at pH 5.1, and VxDE was released from the
resulting membrane fragments by increasing the pH to 7.2 for the final
sonification step. After centrifugation, VxDE was precipitated from the
supernatant by differential
(NH4)2SO4 fractionation and finally
collected by ultracentrifugation (25).
In Vitro De-epoxidation--
For de-epoxidation, VxDE extracts
were diluted 15-fold with 0.4 M citrate-NaOH, pH 5.1. Recombinant LHCII or isolated Vx was mixed with monogalactosyl
diacylglycerol (MGDG) at a molar Vx/MGDG ratio of 1:30 and added to the
assay yielding a final Vx concentration of about 100 nM.
De-epoxidation, performed at 28 °C, was started by the
addition of 30 mM ascorbate. For kinetic
analysis, de-epoxidation was stopped by mixing the sample with
2-butanol at indicated time points. The pigment stoichiometry of
complexes upon incubation with VxDE was determined after concentrating
the sample in 30-kDa Centricon tubes (Amicon), resolubilization in
0.1% n-dodecyl- The pigment composition of LHCII reconstituted with different
carotenoids is summarized in Table I.
2.8-3.0 xanthophylls/monomer were determined in all complexes. In
LHCII reconstituted in the presence of all major xanthophylls the data
were compatible with a stoichiometry of 2 Lu and 1 Nx/monomer, whereas
reconstitution with Vx and Nx in the absence of Lu yielded complexes
containing 1.6 Vx and 1.1 Nx/monomer. Reconstitution of LHCII with Lu
and Vx in the absence of Nx resulted in binding of 2 Lu and about 1 Vx/monomer, and with Vx as the single carotenoid species about 2.8 Vx
were bound per LHCII protein.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helices, have been identified in the crystal
structure and tentatively assigned to Lu (4, 5). Data obtained from
site-directed mutagenesis of LHCII indicated that the Nx binding site
is most likely associated with the more peripheral transmembrane helix
(9). In CP29 (or Lhcb4), one of the minor Chl a/b binding proteins of
PSII, only the two Lu binding sites, called L1 and L2, seem to be
occupied, one by Lu and the other by Vx or Nx (10).
pH-dependent qE-component of NPQ (16), with the
additional requirement of the psbS protein in this process (17). The
xanthophyll cycle pigments are bound to the different antenna
subcomplexes of both photosystems with different stoichiometries
varying from about 0.1 to 1.5/monomer (8, 18-22). Under in
vivo conditions, xanthophyll conversion occurs in various antenna
subcomplexes with different kinetics and to a different extent, which
seems to be related to distinct functions of single subcomplexes in
different components of NPQ (18, 20). It is unclear, however, whether
the differential xanthophyll conversion in single antenna complexes is
determined by specific carotenoid binding sites or simply by a
different accessibility of the Vx de-epoxidase (VxDE) to distinct
antenna complexes. In this work, we investigated the possible role of different carotenoid binding sites for Vx convertibility using the most
abundant Chl a/b protein, LHCII, as a model system for all LHC proteins.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-maltoside and
subsequent purification of monomeric complexes on sucrose gradients.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Pigment stoichiometries of different LHCII complexes
We investigated the convertibility of Vx in the reconstituted
complexes. Under our experimental conditions, free Vx was fully converted to Zx by the VxDE within 10 min (Fig.
1A), with a transient accumulation of the intermediate Ax peaking at 2 min of incubation. A
very similar result was obtained with Lu·Vx monomers containing 2 Lu
and 1 Vx, although the kinetics was retarded by about a factor of 2 (Fig. 1B) when compared with nonbound Vx. In LHCII
containing 1.5 Vx and 1 Nx/monomer, de-epoxidation was even more
strongly retarded. Moreover, the convertibility was restricted to about 50% of the total Vx, indicating a partial accessibility of Vx for
de-epoxidation (Fig. 1C). Replacement of all carotenoids by Vx led to a multiphasic kinetics of de-epoxidation; a fraction of about
60% was convertible to Zx within 30 min as before (Fig. 1,
A and B), whereas the remaining portion
showed a very slow turnover (Fig. 1D).
|
We determined the rate constants for both steps, Vx Ax and Ax
Zx, of the de-epoxidation reactions in all experiments. Because the
de-epoxidation follows a first-order reaction (26-28) and Vx bound to
different sites may exhibit different kinetics, we fitted the data
with 1, 2, or 3 exponentials according to the following scheme.
![]() |
![]() |
|
We further investigated whether the reaction product Zx was re-bound (or stayed bound) to the LHCII complexes. For this purpose, samples were incubated for 70 min under conditions of de-epoxidation. Subsequently, pigmented protein complexes were separated from free pigment by sucrose density gradient centrifugation, and the pigment content of the resulting bands was analyzed by HPLC (summarized in Table III). To distinguish between VxDE-dependent shifts in pigment stoichiometry and potential pigment loss due to incubation temperature (28 °C) and the subsequent reconcentrating step, control samples in the absence of VxDE were also analyzed.
|
The de-epoxidation state (DEPS), defined as Zx/(Zx + Vx), of total pigment samples (i.e. the value shown for free pigment + monomers in Table III) of Nx·Vx-LHCII (0.45) and Vx-LHCII (0.8) after a 70-min VxDE treatment corresponded to the values obtained with the kinetic assays (Fig. 1, C and D). Only Vx·Lu-LHCII (0.9) exhibited a slightly decreased value when compared with the DEPS of 1.0 in the former experiment (Fig. 1B).
Comparing total pigment samples with reisolated monomers, a partial loss of xanthophylls and also of Chl was observed in all samples, including controls, which can be related to the incubation at 28 °C for 70 min. In comparison with untreated monomers (Table I), we observed a loss of about 0.5 Vx in Vx monomers and Vx·Lu complexes leading to 2.3 and 0.5 Vx, respectively. In Nx·Vx complexes, the Vx content remained unchanged whereas Nx was slightly reduced (Table III).
Newly formed Zx was bound to all LHCII complexes with different stoichiometries (Table III). In Vx-LHCII, 1.4 Zx were bound to the monomers while 0.7 Vx withstood the VxDE incubation. The fraction of VxDE-resistant Vx was even greater in Nx·Vx-LHCII, where 1.2 Vx and only 0.3 Zx were bound. The remaining 0.5 Vx in the control samples of Vx·Lu monomers was almost completely converted to 0.4 Zx with only minor amounts of Vx persisting VxDE treatment. Thus, somewhat reduced if similar DEPS values were found in the reisolated monomers as noted before in the kinetic analysis, indicating that the newly formed Zx molecules were rebound by or stayed bound to the complexes.
De-epoxidation with Vx·Nx-LHCII containing 1.6 Vx and nearly 1 Nx/monomer (Fig. 1C) resulted in a maximum DEPS of about 0.5, indicating that about 1 Vx/monomer was inaccessible for de-epoxidation. This partial conversion to Zx could result from two different restrictions. Assuming that the Vx molecules are bound to the two Lu binding sites, L1 and L2, of the native complex, de-epoxidation could be generally suppressed if one of the two Lu binding sites is occupied by Zx independently of the binding site. Alternatively, one specific binding site, either L1 or L2, could be unavailable for de-epoxidation, e.g. for structural reasons.
We designed the following experimental approach to decide between these
two possibilities. LHCII complexes were reconstituted with a mixture of
Nx, Vx, and Zx yielding stoichiometries of 1 Nx/momomer in all cases
but with Vx/Zx ratios varying between 1.9:0.5 and 0.03:1.9/monomer
(Fig. 2A). Fig. 2B
indicates an equivalent relative binding affinity
KVx/Zx of 0.24 in L1 and L2. The absence of any
biphasic behavior of the competition curve with a slope of 0.24 strongly suggests that L1 and L2 preferentially bind Zx with respect to
Vx but do not differ in their relative affinities toward these
carotenoids.
|
We expected that no de-epoxidation would occur if the presence
of Zx in one of the two Vx binding sites generally restricts de-epoxidation of the Vx present in these complexes. This should become
particularly obvious in complex preparations with a Zx/Vx ratio 1. On the other hand, de-epoxidation of about 50% of the Vx should be
observed again, if one specific binding site is inaccessible for
de-epoxidation and if both binding sites have a similar affinity toward
Vx and Zx. In fact, at all Vx/Zx ratios we found that de-epoxidation was limited to about 50% of the respective Vx pool. This is shown as
an example for a Vx/Zx ratio of 1:1 in Fig.
3. The rate constants for the reactions
were in a similar range as in the respective experiments with Vx·Nx
monomers (Table II). The observation that only 50% of Vx is
convertible to Zx, irrespective of the Zx/Vx ratio in the reconstituted
complexes, indicates that Vx is accessible to the VxDE only in one of
the two Lu binding sites (L1 or L2).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We have shown for the first time that Vx bound to solubilized LHCII can be de-epoxidized in vitro by partially purified VxDE prepared from spinach thylakoids. It has been found2 that purified VxDE did not de-epoxidize Vx bound to pigment protein fractions of LHCII even when the latter was supplemented with MGDG (19). In these experiments, it is most likely trimeric LHCII isolated from thylakoid membranes was used, whereas our data were obtained with recombinant monomeric LHCII. In experiments with the recombinant trimeric forms of the different complexes, however, we have also observed de-epoxidation, but to a lower extent (maximum 50% conversion) and at much slower rates (a factor of about 10; data not documented). Thus, it is reasonable to assume that Vx bound to trimeric LHCII is not readily accessible for in vitro de-epoxidation. Because our experiments were performed at 28 °C, Vx might have become accessible by thermally induced destabilization of the trimers.
Incubation of complexes under de-epoxidation conditions and subsequent re-purification leads to a preferential loss of Chl a (compare Table I and control monomers in Table III). The normalization of initially isolated versions of LHCIIb to 13 Chls and reisolated monomers to 12 Chls is justified by this loss. Otherwise, if a number of 12 Chls would be assumed in the original samples, the insertion of up to 1 Chl b after treatment for de-epoxidation had to be considered, which is very unlikely.
Our pigment data indicate binding of almost 3 carotenoids/monomer in all complexes, in rough agreement with previous studies (11, 12). For LHCII containing either Vx as its single carotenoid species or Lu plus Vx, lower stoichiometries of 2.2-2.3 have also been reported (11, 12). These differences are most likely based on varying purification conditions and reflect a low affinity of Vx to the Nx binding site of the native protein.
According to previous work (11, 12), we assume that in complexes containing 2 Lu and 1 Vx, the two Lu binding sites of the native complex, L1 and L2, are both occupied by Lu and that Vx is bound either to the Nx binding site of the native complex or to a peripheral site. The notion that Vx is most rapidly converted to Zx in complexes containing 2 Lu and 1 Vx could then be explained by assuming that Vx is easily accessible for VxDE at these sites.
In complexes reconstituted with Vx and Nx, 1.6 Vx and 1 Nx were determined per monomer. Because Nx has a much higher affinity to the Nx binding site of the native complex in comparison with Vx (11), it is likely that in these complexes the Vx molecules are bound to the L1 and L2 site of LHCII. Our experiments have shown that only a portion of this Vx can be converted into Zx (Fig. 1C) and that Vx bound to one specific site, either L1 or L2, is not available for de-epoxidation (Fig. 3). This may indicate that one of the two Lu binding sites essentially requires binding of a carotenoid, whereas the other one could, at least transiently, be empty. A similar conclusion has been drawn from recent studies with recombinant Lhcb4 (CP29) (10). In this monomeric antenna protein of PSII, the L1 site is occupied by Lu, whereas the L2 site can bind either Vx or Nx (10). Vx bound to Lhcb4 is known to be de-epoxidized in vivo under high light stress (18, 20). Thus the proposed structural similarity among all Chl a/b-binding proteins would support the assumption that Vx bound to the site L2 of LHCII can be de-epoxidized to Zx, whereas the Vx bound to site L1 cannot. This is further in accordance with the observation that the L1 site is obligatory occupied by Lu in all Chl a/b-binding proteins analyzed so far, whereas the L2 site can be occupied by Vx to various extents in the different complexes (8).
In LHCII complexes containing Vx as the single carotenoid species, all three binding sites are occupied by Vx. In contrast to the Vx·Nx monomers, Vx bound to both the L1 and L2 site was convertible to Zx. Obviously, the unexpected accessibility of Vx bound to L1 was related to the absence of Nx in these complexes. This interpretation would suggest a stabilizing function of Nx for the structure of LHCII. Because we did not observe a pronounced dissociation of these complexes, the lack of Nx may simply result in a slightly changed overall conformation providing an increased accessibility of Vx bound to L1. Two of the three Vx were rapidly convertible to Zx in these complexes with similar kinetics as found in the Lu·Vx and Vx·Nx monomers (Fig. 1, Table II). This finding confirms the conclusion that Vx bound to the L2 site and the Nx binding site of the native complex is easily accessible for de-epoxidation.
Our analyses of the reaction kinetics indicated that Vx is
de-epoxidized at different carotenoid binding sites at specific rates
(Table II). It cannot be decided from our data whether this specificity
is determined by different binding affinities of Vx to each site or by
a different accessibility of VxDE to the three carotenoid binding
sites. With the exception of the slowest phase, determined in the
experiment with Vx monomers (about 0.01 min 1, Table II), the rate constants for
de-epoxidation are in agreement with the kinetics found in earlier
studies at saturating light intensities in intact leaves (26, 27) or,
under optimum conditions, in isolated thylakoids (28, 29). Thus, it is
reasonable to assume that the kinetics determined with recombinant
LHCII and VxDE extracts can be applied also to membrane bound antenna proteins.
It is known that most of Vx present in thylakoid membranes is bound to PSI antenna proteins and to the minor PSII antenna proteins Lhcb4-6 but not to LHCII (8, 18-20). According to the structural similarity among the different antenna proteins, the characteristics of Vx de-epoxidation at recombinant LHCII may be used as a model for all Chl a/b-binding proteins in higher plants. Furthermore, the analysis of Lu-deficient mutants of Arabidopsis have shown that Lu can presumably be replaced even under in vivo conditions by other xanthophylls, in particular Vx and Ax, without affecting the viability of the plants (30, 31).
The kinetic analyses and the fact that the reaction product Zx is
(re)bound to the LHC II complexes further provide new insights into the
mechanism of de-epoxidation. The kinetic analyses have shown that, in
contrast to the first step (Vx Ax) of de-epoxidation, the kinetics
of the second Ax
Zx step are nearly identical in all experiments
and thus independent of the Vx binding site. This leads to ratios of
k2/k1(1,2,3) varying from
2 (in Vx·Lu monomers) to 80 (slowest phase in Vx monomers) indicating
that both reactions are rate-limited by different factors. According to
our data, the first step (Vx
Ax) seems to be kinetically controlled
by the binding strength of Vx to the LHCII complex (or the
accessibility of the VxDE to the binding site), whereas the Ax
Zx
step may rather be rate-limited by processes related to the VxDE
itself. It is tempting to speculate that the dynamic interaction of the intermediate Ax with VxDE (which is required to make the second epoxy
group accessible to the active center of the enzyme) determines the
kinetics of the second step. Because the Ax
Zx conversion follows
similar kinetics in the presence and absence of LHCII complexes, the
movement of Ax must be independent of antenna proteins. This
lets us speculate that the intermediate Ax is not bound to the LHC II
complexes but rather is associated with the VxDE. After de-epoxidation
of the second epoxy group, the product Zx can be rebound by the LHCII complexes.
It is unknown to what extent xanthophylls are released into the lipid phase after de-epoxidation under in vivo conditions. At least for LHCII and Lhcb4 it has been reported that occupation of some of the binding sites depends on the de-epoxidation state of the xanthophyll cycle pigments (20, 21). Both a redistribution of xanthophyll cycle pigments among chlorophyll-binding proteins (20) as well as a partial release into the lipid phase (21) can be assumed on the basis of these studies. Moreover, recent analyses of xanthophyll cycle mutants from Arabidopsis thaliana indicated that xanthophylls may serve important functions not only in energy dissipation but also as membrane stabilizers and antioxidants in the lipid phase (32, 33). The latter characteristics resemble the proposed functions of tocopherols in the membrane (34). Thus, at least a partial release of Zx into the lipid phase would make sense as judged from the physiological functions of xanthophylls.
In our experiments with recombinant LHCII, however, the xanthophyll/Chl ratios of the de-epoxidized and control LHCII monomers were only slightly different in Vx complexes (0.18 and 0.19, respectively) and were identical for Nx·Vx-LHCII and Vx·Lu-LHCII, although Zx was also present in the free pigment zone (Table III). Obviously, the partial loss of xanthophylls was independent of the DEPS and was caused simply by the incubation at 28 °C. We conclude from this result that under our experimental conditions accessible Vx is converted to Zx and, provided that complexes do not fall apart, all newly formed Zx is rebound by the LHCII complexes. Considering the low concentrations of LHCII complexes and carotenoids in our assay, it can be assumed that LHCII and VxDE stay in close contact during the de-epoxidation reactions and that no release of carotenoids into the lipid phase occurs.
In conclusion, our data support the view that the kinetics and the
extent of Vx de-epoxidation is controlled by the carotenoid binding
site of a distinct antenna protein. Thus, it is very likely that the
differential xanthophyll conversion in single antenna complexes
observed under in vivo conditions (18, 20) is determined by
the characteristics of the Vx binding sites and not by the accessibility of VxDE to the antenna proteins. Further work is now
needed to characterize the Vx de-epoxidation in other antenna proteins
and to explore the influence of the xanthophyll conversion of
protein modifications that occur under different light conditions (binding of protons, phosphorylation) and that are likely to be central
to the regulation of light harvesting.
![]() |
ACKNOWLEDGEMENTS |
---|
The help of S. Raunser with the preparation of some of the reconstituted samples is gratefully acknowledged. Zx was a kind gift from Roche Diagnostics (Switzerland).
![]() |
FOOTNOTES |
---|
* This work was supported by the Deutsche Forschungsgemeinschaft SFB 189, TP B13, and Ja 665/2-1 (to P. J.) and Pa-324/5-3 (to H. P.) and a grant from the Stiftung Rheinland Pfalz für Innovation (to H. P.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ To whom correspondence should be addressed. Tel.: +49-211-81-13862; Fax: +49-211-81-13706; E-mail: pjahns@uni-duesseldorf.de.
Published, JBC Papers in Press, April 11, 2001, DOI 10.1074/jbc.M102147200
2 K. Hindehoffer, A. Lee, P. Thornber, and H. Y. Yamamoto, unpublished work.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: Chl, chlorophyll; Ax, antheraxanthin; DEPS, de-epoxidation state; HPLC, high pressure liquid chromatography; LHC, light-harvesting complex; Lu, lutein; MGDG, monogalactosyl diacylglycerol; NPQ, non-photochemical quenching of chlorophyll fluorescence; Nx, neoxanthin; PSI, photosystem I; PSII, photosystem II; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; Vx, violaxanthin; VxDE, violaxanthin de-epoxidase; Zx, zeaxanthin.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Green, B. R., Pichersky, E., and Kloppstech, K. (1991) Trends Biochem. Sci. 16, 181-186[CrossRef][Medline] [Order article via Infotrieve] |
2. | Jansson, S. (1994) Biochim. Biophys. Acta 1184, 1-19[Medline] [Order article via Infotrieve] |
3. | Jansson, S. (1999) Trends Plant Sci. 4, 237-240 |
4. | Kühlbrandt, W., and Wang, D. N. (1991) Nature 350, 130-134[CrossRef][Medline] [Order article via Infotrieve] |
5. | Kühlbrandt, W., Wang, D. N., and Fujiyoshi, Y. (1994) Nature 367, 614-621[CrossRef][Medline] [Order article via Infotrieve] |
6. |
Peter, G. F.,
and Thornber, J. P.
(1991)
J. Biol. Chem.
266,
16745-16754 |
7. | Bassi, R., Pineau, B., Dainese, P., and Marquardt, J. (1993) Eur. J. Biochem. 212, 297-303[Abstract] |
8. | Bassi, R., and Caffarri, S. (2000) Photosynth. Res. 64, 243-256[CrossRef] |
9. | Croce, R., Remelli, R., Varotto, C., Breton, J., and Bassi, R. (1999) FEBS Lett. 456, 1-6[CrossRef][Medline] [Order article via Infotrieve] |
10. | Bassi, R., Croce, R., Cugini, D., and Sandona, D. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 0056-10061 |
11. |
Croce, R.,
Weiss, S.,
and Bassi, R.
(1999)
J. Biol. Chem.
274,
29613-29623 |
12. |
Hobe, S.,
Niemeier, H.,
Bender, A.,
and Paulsen, H.
(2000)
Eur. J. Biochem.
267,
616-624 |
13. | Demmig-Adams, B., and Adams, W. W., III (1992) Annu. Rev. Plant Physiol. Plant Mol. Biol. 43, 599-626[CrossRef] |
14. | Horton, P., Ruban, A. V., and Walters, R. G. (1996) Annu. Rev. Plant Physiol. Plant Mol. Biol. 47, 655-684[CrossRef] |
15. | Niyogi, K. K. (1999) Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 333-359[CrossRef] |
16. |
Niyogi, K. K.,
Grossman, A. R.,
and Björkman, O.
(1998)
Plant Cell
10,
1121-1134 |
17. | Li, X. P., Björkman, O., Shih, C., Grossman, A. R., Rosenquist, M,., Jansson, S., and Niyogi, K. K. (2000) Nature 403, 391-395[CrossRef][Medline] [Order article via Infotrieve] |
18. |
Färber, A.,
Young, A. J.,
Ruban, A. V.,
Horton, P.,
and Jahns, P.
(1997)
Plant Physiol.
115,
1609-1618 |
19. | Yamamoto, H. Y., and Bassi, R. (1996) in Oxygenic Photosynthesis: The Light Reactions (Ort, D. R. , and Yocum, C. F., eds) , pp. 539-563, Kluwer Academic Publishers, Dordrecht, The Netherlands |
20. |
Verhoeven, A. S.,
Adams, W. W., III,
Demmig-Adams, B.,
Croce, R.,
and Bassi, R.
(1999)
Plant Physiol.
120,
727-737 |
21. |
Ruban, A. V.,
Lee, P. J.,
Wentworth, M.,
Young, A. J.,
and Horton, P.
(1999)
J. Biol. Chem.
274,
10458-10465 |
22. |
Schmid, V. H. R.,
Cammarata, K. V.,
Bruns, B. U.,
and Schmidt, G. W.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
7667-7672 |
23. | Paulsen, H., Finkenzeller, B., and Kuehlein, N. (1993) Eur. J. Biochem. 215, 809-816[Abstract] |
24. | Martinson, T. A., and Plumley, F. G. (1995) Anal. Biochem. 228, 123-130[CrossRef][Medline] [Order article via Infotrieve] |
25. | Arvidsson, P.-O, Bratt, C. E., Carlsson, M., and Åkerlund, H.-E. (1996) Photosynth. Res. 49, 119-129 |
26. |
Härtel, H.,
Lokstein, H.,
Grimm, B.,
and Rank, B.
(1996)
Plant Physiol.
110,
471-482 |
27. |
Jahns, P.
(1995)
Plant Physiol.
108,
149-156 |
28. | Siefermann, D., and Yamamoto, H. Y. (1974) Biochim. Biophys. Acta 357, 144-150[Medline] [Order article via Infotrieve] |
29. |
Pfündel, E.,
and Dilley, R. A.
(1993)
Plant Physiol.
101,
65-71 |
30. |
Pogson, B.,
McDonald, K. A.,
Truong, M.,
Britton, G.,
and DellaPenna, D.
(1996)
Plant Cell
8,
1627-1639 |
31. |
Pogson, B.,
Niyogi, K. K.,
Björkman, O.,
and DellaPenna, D.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
13324-13329 |
32. |
Havaux, M.,
and Niyogi, K. K.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
8762-8767 |
33. |
Havaux, M.,
Bonfils, J.-P.,
Lütz, C.,
and Niyogi, K. K.
(2000)
Plant Physiol.
124,
273-284 |
34. | Fryer, M. (1992) Plant Cell Environ. 15, 381-392 |