JBC Focus on PI3-Kinase with Echelon

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M402461200 on March 21, 2004

J. Biol. Chem., Vol. 279, Issue 22, 22866-22874, May 28, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/22/22866    most recent
M402461200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Li, X.-P.
Right arrow Articles by Niyogi, K. K.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Li, X.-P.
Right arrow Articles by Niyogi, K. K.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Regulation of Photosynthetic Light Harvesting Involves Intrathylakoid Lumen pH Sensing by the PsbS Protein*

Xiao-Ping Li{ddagger}§, Adam M. Gilmore§||, Stefano Caffarri**{ddagger}{ddagger}§§, Roberto Bassi**{ddagger}{ddagger}, Talila Golan{ddagger}, David Kramer¶¶, and Krishna K. Niyogi{ddagger}||||

From the {ddagger}Department of Plant and Microbial Biology, University of California, Berkeley, California 94720-3102, the Ecosystem Dynamics Group, Research School of Biological Sciences, Institute of Advanced Studies, Australian National University, Canberra, ACT 2601, Australia, the **Université Aix-Marseille II, Laboratoire de Genetique et Biophysique des Plants, Département de Biologie-Case 901, 163 Avenue de Luminy-13288, Marseille, France, the {ddagger}{ddagger}Dipartimento Scientifico e Tecnologico, Università di Verona, Strada Le Grazie 15, 37134, Verona, Italy, and the ¶¶Institute of Biological Chemistry and Department of Biochemistry and Biophysics, Washington State University, Pullman, Washington 99164-6340

Received for publication, March 4, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The biochemical, biophysical, and physiological properties of the PsbS protein were studied in relation to mutations of two symmetry-related, lumen-exposed glutamate residues, Glu-122 and Glu-226. These two glutamates are targets for protonation during lumen acidification in excess light. Mutation of PsbS did not affect xanthophyll cycle pigment conversion or pool size. Plants containing PsbS mutations of both glutamates did not have any rapidly inducible nonphotochemical quenching (qE) and had similar chlorophyll fluorescence lifetime components as npq4-1, a psbS deletion mutant. The double mutant also lacked a characteristic leaf absorbance change at 535 nm ({Delta}A535), and PsbS from these plants did not bind dicyclohexylcarbodiimide (DCCD), a known inhibitor of qE. Mutation of only one of the glutamates had intermediate effects on qE, chlorophyll fluorescence lifetime component amplitudes, DCCD binding, and {Delta}A535. Little if any differences were observed comparing the two single mutants, suggesting that the glutamates are chemically and functionally equivalent. Based on these results a bifacial model for the functional interaction of PsbS with photosystem II is proposed. Furthermore, based on the extent of qE inhibition in the mutants, photochemical and nonphotochemical quenching processes of photosystem II were associated with distinct chlorophyll fluorescence life-time distribution components.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In conditions of excess light, photosynthetic light harvesting is regulated by a feedback de-excitation mechanism termed energy-dependent quenching (qE),1 which increases thermal dissipation of excess absorbed light energy in photosystem II (PSII). The qE mechanism is triggered by conditions that limit photosynthetic carbon fixation and result in increased acidification of the chloroplast thylakoid lumen (1–4). The thermal dissipation of excess excitation energy is most commonly measured and referred to as nonphotochemical quenching (NPQ) of PSII chlorophyll (Chl) a fluorescence. Although there are several components of NPQ, in higher plants qE can account for the major part of NPQ and is characterized by its relatively fast induction and relaxation kinetics, on a physiological time scale of seconds to minutes. The decrease in the intensity of Chl fluorescence is the result of the decrease in the electronic excited state lifetime of Chl caused by an increased thermal dissipation rate constant (5).

The rapid response of the qE process is chemically associated with changes in the trans-thylakoid membrane pH gradient ({Delta}pH). The {Delta}pH change has at least two functions in qE. First, it activates the violaxanthin de-epoxidase that converts violaxanthin (V) to antheraxanthin (A) and zeaxanthin (Z) (6). A and/or Z are essential elements of qE (7–9). Second, the lower pH in the lumen results in protonation of PSII proteins, including the 22-kDa PSII subunit, PsbS, which plays a key role in qE (10). When both pH-induced changes occur together it is believed that Chls in PSII can transfer their excess energy to Z, which can return to the ground state via thermal decay (7, 11, 12).

The pH-sensing mechanism of the PsbS protein is influenced by two pairs of symmetrically arranged glutamate residues, each located within or close to the two lumen-exposed loops of the protein (13). Dicyclohexylcarbodiimide (DCCD), a well known inhibitor of qE (14–16) is a carboxylate-modifying agent (17) that binds to PsbS (18). Although it was suggested that the DCCD binding site is in the lumenal loops of PsbS, the exact binding site has not been determined. Importantly, site-directed mutagenesis experiments indicated that two of the PsbS glutamates, Glu-122 and Glu-226, are necessary for the function of PsbS (13).

In this article we used single and double mutations of PsbS (E122Q/E226Q) to make a detailed biochemical and biophysical analysis of the role of these two glutamates in pH sensing and DCCD binding. We probed the role of the Glu-122 and Glu-226 residues by monitoring the changes in the PSII Chl a fluorescence lifetime distributions, intensities, and photochemical efficiencies. Additionally, we also monitored the absorbance change at 535 nm ({Delta}A535) (19) that is obligatorily associated with qE (10, 20, 21). The {Delta}A535 has been suggested to reflect a red shift in the absorption spectrum of Z that occurs upon binding to PsbS (22, 23). From these measurements we formulated a model of the influence of the glutamate mutants on the fractional quenching of the populations of PSII (24) that is consistent with the symmetrical structure of the PsbS protein, the DCCD binding stoichiometry, and the stoichiometry of the xanthophyll components on a per PSII unit basis.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Plant Materials and Growth Conditions—Arabidopsis thaliana (Columbia strain) was used in all experiments. npq4-1 is a psbS deletion mutant (10). Site-directed mutagenesis of psbS and transformation protocols were the same as described previously (13). In the T1 generation, lines with single insertions were selected, and homozygous lines were confirmed in the T3 generation. All plants were grown in a growth chamber with a light intensity of 120 µmol photons m–2 s–1 and a photoperiod of 10 h light (22 °C)/14 h dark (20 °C).

DNA and Immunoblotting—psbS DNA gel blotting and PsbS protein immunoblotting analyses were performed as described previously (13), except that genomic DNA was digested with SpeI.

NPQ Measurement—Room temperature Chl fluorescence was measured on the attached rosette leaves with a commercial fluorometer (FMS2, Hansatech, King's Lynn, United Kingdom). Plants were dark-adapted overnight before the measurement. After measuring Fm during a 1.0-s saturating pulse of 10,000 µmol photons m–2 s–1, actinic light of 1200 µmol photons m–2 s–1 was switched on for 10 min. A saturating 1.0-s pulse of 10,000 µmol photons m–2 s–1 was triggered every 60 s to determine . NPQ was calculated as .

Cross-linking and Immunoblot Analysis—Fresh thylakoids were extracted from mature rosette leaves (24). Thylakoids containing 50 nmol of Chl were diluted in 1 ml of reaction buffer (50 mM HEPES, pH 8, 0.1 M sucrose, 10 mM NaCl, 10 mM KCl, 5 mM MgCl2, 1mM KH2PO4, 30mM sodium ascorbate, 50 µM methyl viologen, 0.3 mM ATP) and stirred in the dark or under qE-inducing illumination. After 10 min, dithiobis(succinimidylpropionate) dissolved in dimethyl sulfoxide was added to a final concentration of 0.1 mM and stirred for 10 min. Cross-linking was terminated with the addition of Tris-HCl (pH 8) to 20 mM and stirring for an additional 10 min. The cross-linked thylakoids were centrifuged for 2 min at 15,300 x g and resuspended with protein solubilization buffer without reductant for analysis by SDS-PAGE. For immunoblotting of cross-linked samples, 5 nmol of Chl was loaded in each lane.

Xanthophyll Cycle Pigment Analysis—For zeaxanthin formation experiments, leaf discs were sampled from overnight dark-adapted plants as controls, sampled at noon in the growth chamber (low light conditions), and sampled at noon, floated on water, and then treated at 1600 µmol photons m–2 s–1 for 30 min (high light-treated). All samples were rapidly frozen in liquid nitrogen and stored at –80 °C until analysis. Pigments were measured by high performance liquid chromatography (25), and the mean ± S.D. was calculated (n = 6–9).

Leaf Absorbance Measurements—Leaf absorbance changes were measured using the Non-Focusing Optics Spectrophotometer (NoFOSpec) (26), but modified as described (27) to allow semi-simultaneous measurements of absorbance changes at four different wavelengths. This was accomplished by aiming four separate banks of light-emitting diodes (HLMP-CM15, Agilent Technologies, Santa Clara, CA), each filtered through a separate 5-nm band-pass interference filter (Omega Optical, Brattleboro, VT), into the entrance of a compound parabolic concentrator. Each bank of light-emitting diodes was filtered with a separate interference filter, at 500, 520, 535, and 545 nm each with a 5-nm band pass (full width at half-height). The photodiode detector was protected from direct actinic light by a Schott BG-18 filter. Current from the photodiode was converted to a voltage by an operational amplifier, and the resulting signal was AC-filtered to remove background signals, and sampled by a 16-bit analog-to-digital converter on a personal computer data acquisition card (DAS16/16-AO, Measurement Computing, Middleboro, MA). Timing pulses were generated by digital circuitry (PC card D24/CTR 3, Measurement Computing, Middleboro, MA) controlled by software developed in-house. The duration of the probe pulses was set at 10 µs. Actinic illumination was provided by a set of 12 red light-emitting diodes (HLMP-EG08-X1000, Agilent Technologies) and controlled by the timing circuitry. Measuring pulses at each wavelength were given in sequence at 1–100-ms intervals, depending upon the experiment. Chl a fluorescence changes were also measured with the NoFOSpec instrument using the 525-nm measuring pulse to excite Chls, while protecting the detector with a Schott RG-9 filter (28).

Leaves were cut from plants dark-adapted overnight, and the leaf petiole was wrapped in a small piece of moist cotton. The Fm was first recorded by giving 800 ms white light of 30,000 µmol photons m–2 s–1. After 30 s in the dark, a red actinic light of 1300 µmol photons m–2 s–1 was switched on for 215 s, and the leaf absorption at 535 nm was recorded. After allowing relaxation of qE in the dark for 110 s, leaf absorption at 535 nm was recorded again, and the {Delta}A535 was calculated. After finishing the 535-nm measurement, the same actinic light was switched on again for 2 min, and then a white light pulse (800 ms, 30,000 µmol photons m–2 s–1) was applied to measure the .

DCCD Binding Analysis—[14C]DCCD labeling was conducted on thylakoids prepared as described previously (29) at two different pH conditions, pH 7.8 and 5, for 3 h at room temperature. For labeling at pH 7.8, thylakoid samples containing 30 µg of Chl were labeled in buffer T2 (5 mM Tricine, pH 7.8, 50 mM sorbitol, 10 mM EDTA). For labeling at pH 5, thylakoid samples were labeled in citrate buffer (30 mM sodium citrate, pH 5, 50 mM sorbitol, 10 mM EDTA). After mixing thylakoids with buffer, 7.5 µlof100 µM [14C]DCCD (0.5 µCi) in ethanol was added to make a final total volume of 100 µl. For gel analysis, a first dimension separation using a 12% acrylamide Tris sulfate gel with 6 M urea was used (30). After staining the gel with Coomassie Blue, the region corresponding to PsbS and light-harvesting complex proteins was cut out and equilibrated in Tris-HCl buffer, pH 6.8, with 2% (w/v) SDS for 25 min, then transferred to the same buffer containing 60% glycerol for another 10 min. The gel slice was then loaded for second dimension separation by SDS-PAGE on a 10–16% gel without urea (31). The labeling intensities of PsbS mutants with respect to the wild type were acquired from the dried gels with the Instant Imager (Packard Instrument Co.) instrument and calculated from the average of three independent measurements after correcting for the quantity of PsbS on the SDS-PAGE.

Room Temperature Chl Fluorescence Lifetime Measurement—Leaf samples were dark-adapted for 12 h at room temperature prior to detachment at the petiole and complete vacuum infiltration of the intercellular airspace with 0.35 M glucose (30 min in dimmed room light at room temperature) in a 5-ml disposable syringe. Infiltrated leaves were gently pressed using cotton wool into a front surface configuration free of air bubbles in a low-fluorescence plastic-quartz cuvette submerged in 0.35 M glucose solution. The samples were placed in an aluminum block-type sample holder that could be temperature controlled using circulating water baths. After dark adaptation for 10 min at 20 °C the samples were illuminated, and Chl a fluorescence was monitored using the fiber-optic probe of a Chl fluorometer (PAM 101–103, Heinz-Walz Instruments Ltd., Effeltrich, Germany). Experiments were initiated at 20 °C with the low-intensity modulated measuring beam (1.6 KHz, {approx}0.25 µmol photons m–2 s–1, 440 nm) to measure the PSII fluorescence intensity (interference filter 685 nm, 10-nm bandpass) under conditions of maximal photochemistry (Fo). After 30 s Fm was measured during a saturating 2-s pulse of white light (13,600 µmol photons m–2 s–1, Walz DT-Cyan filter), and then a strong actinic white light (1250 µmol photons m–2 s–1, Walz DT-Cyan filter) was applied to induce photosynthetic electron transport, thylakoid lumen acidification, V de-epoxidation, and PsbS protonation. Saturating pulses were applied every 60 s to measure . After 10 min the sample temperature was reduced to 3 °C (requiring ~3 min after switching water baths), whereas the actinic light was continued for a total of 15 min. All illumination was extinguished, and the sample was then rapidly turned to face the fiber-optic probe of the fluorescence lifetime instrument to illuminate the sample (140 µmol photons m–2 s–1, 635 nm) for 5 min before initiating the fluorescence lifetime determination. Fluorescence lifetime determinations were made under conditions of light- and low temperature (3 °C)-saturated lumen acidification (32) to maintain maximal levels of V de-epoxidation and PsbS protonation, whereas the PSII redox state was ~60–80% reduced, depending on the level of energy dissipation (33). The fluorescence intensity conditions during the lifetime acquisition corresponded to those defined as Fs (34).

The PSII Chl a fluorescence lifetimes were determined using a multifrequency phase-modulation fluorimeter (model K2–004, ISS Instruments, Urbana, IL) using a red laser diode (peak emission 635 nm) for excitation and a red-sensitive microchannel plate photomultiplier tube (Hamamatsu Photonics, Hamamatsu, Japan; R3809U-50) for emission detection. Excitation and emission were applied and measured, respectively, from the front surface of the sample using a bifurcated quartz fiber-optic probe with the single terminus facing the sample. The excitation diode-laser intensity was attenuated to 140 µmol photons m–2 s–1 at 30 MHz to avoid photobleaching during measurements, which normally required 10 min to complete. The reference and sample signals were detected at each modulation frequency using high transmittance (>80%) narrow waveband (12 nm half-width) interference filters (Corion Inc., Franklin, MA) centered at 645 and 689 nm, respectively. An automatic rotating filterwheel exchanged the sample and reference positions such that the phase and modulation signals were repeatedly determined using 4–6 cycles at each modulation frequency until standard errors were reduced below 0.04 degrees for the phase angle shift and 0.001 for the demodulation ratios.

Global Analysis of the Phase and Modulation Fluorescence Lifetime Data—The phase angle shift and demodulation ratio data sets for all samples were analyzed globally by fitting to a multimodal Lorentzian distribution model with the {tau}-center value and width of each fractional component being linked together and all amplitudes free-floating. Similar to the analysis described previously (33), a minimum of five positive and one negative amplitude components were necessary to fit the data from all sample types globally. The goodness of fit for the data set, which included 62 free fitting parameters and 404 data points, was judged on the following criteria: the residual error distribution was randomly and normally distributed around 0 after minimizing the reduced {chi}-square to a value of 2.0223 using frequency-independent standard deviation values {sigma}phase shift = 0.25° and {sigma}demodulation = 0.005 (35). Global analysis was performed on a program assembled with Excel 2002, Visual Basic (Microsoft) and Large-Scale GRG Solver Engine (Frontline Systems, Inc., Incline Village, NV). The program was evaluated by using statistical reference datasets provided by the National Institute of Standards and Technology.2


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Effect of Glutamate Mutations on PsbS Function and qE— Glutamate residues in the lumenal loops of PsbS were changed to glutamines by site-directed mutagenesis to eliminate H+-binding capacity while minimizing alteration of the protein structure (13). The E122Q and E226Q mutations were made individually and pairwise and expressed in npq4-1 Arabidopsis plants that lack endogenous wild-type PsbS. As a control, npq4-1 plants were transformed with the wild-type psbS gene. For each single mutant (npq4-E122Q and npq4-E226Q), the double mutant (npq4-E122Q,E226Q), and the control (npq4-1 + psbS), two to three independent lines carrying single insertions were selected (Fig. 1A).



View larger version (66K):
[in this window]
[in a new window]
  		FIG. 1.
DNA gel blot analysis and protein immunoblot analysis of Arabidopsis wild-type and PsbS mutant plants. Shown here are the wild-type, three lines of npq4-1 + psbS, three E122Q mutant lines (npq4-E122Q), two E226Q mutant lines (npq4-E226Q), two double mutant lines (npq4-E122Q,E226Q), and npq4-1. For DNA gel blotting 10 µg of genomic DNA was digested with SpeI prior to loading. For immunoblots 1.0 nmol of Chl was loaded in each lane, and PsbS and D1 proteins were detected using specific antibodies.

 
Immunoblot analysis showed that PsbS protein levels in all transgenic lines were 3–5-fold higher than in wild-type plants (Fig. 1). Immunoblotting was conducted on washed thylakoid membranes, demonstrating that the PsbS protein was inserted in the membrane in all cases. Furthermore, protein cross-linking experiments with isolated thylakoids showed no differences in PsbS cross-linking patterns (data not shown), indicating that the glutamate mutations of PsbS and enhanced protein expression level do not affect the physical association of PsbS within the thylakoid membranes or its interaction with other proteins.

Fig. 2 shows the NPQ induction traces in the transgenic lines, the wild type, and npq4-1. Compared with the npq4-1 + psbS lines, which had higher levels of NPQ than the wild type because of overexpression of PsbS (33), the single mutant lines had about one-third as much NPQ. Most of the NPQ was rapidly reversible (qE). There was no significant difference between the single mutant lines npq4-E122Q and npq4-E226Q. The NPQ level in the npq4-E122Q,E226Q double mutant lines was indistinguishable from that of npq4-1, which lacks qE, indicating that the double mutant PsbS was nonfunctional.



View larger version (28K):
[in this window]
[in a new window]
  		FIG. 2.
Induction and relaxation of NPQ. Measurements were conducted on two separate batches of dark-adapted plants with 3 to 5 replications each. NPQ was measured during 10 min of illumination with high light (1200 µmol photons m–2 s–1), followed by relaxation in the dark for 5 min. All data points were plotted as the mean ± S.E. (n = 8).

 
Xanthophyll Cycle Pigment Composition and {Delta}A535—There were no significant differences in either the total xanthophyll cycle pigment pool sizes or V de-epoxidation states in high light (1700 µmol photons m–2 s–1) in any of the different lines sampled at the same conditions (data not shown). The xanthophyll cycle pool size was 26.5 ± 0.8, 28.2 ± 0.9, and 29.6 ± 1.2 mmol/mol Chl a in the three sample conditions, and the respective de-epoxidation states were 0.053 ± 0.010 in overnight dark-adapted leaves, 0.068 ± 0.010 in leaves sampled at noon under growth light conditions, and 0.649 ± 0.010 in leaves treated for 30 min with light that was 10 times higher than the growth light. Thus, we conclude that neither the amount of PsbS in the thylakoid membrane, as shown before (33, 36), nor the mutations of the PsbS protein influence the xanthophyll cycle pool size or de-epoxidation in high light.

Fig. 3A shows that {Delta}A535 was linearly correlated with the extent of NPQ in each sample. There was a significant x intercept value of about 0.36, indicating that a small component of the total NPQ is independent of {Delta}A535 and PsbS. This component is attributable to qI, a component of NPQ that is related to photoinhibitory damage to PSII and associated with a slowly reversing component of Fm quenching that is observed in the absence of PsbS, as discussed in more detail below. We further analyzed the linear relationship between {Delta}A535 and NPQ by consolidating the variations of the sampling in each measurement and plotting the lines for the mean values of NPQ and {Delta}A535 from the wild type, npq4-1, npq4-1 + psbS, npq4-E122Q, npq4-E226Q, and npq4-E122Q,E226Q lines (Fig. 3B). This analysis increased the significance of the slope and intercept values (Table I). There were no significant differences between npq4-1 and the double mutant or between the npq4-E122Q and npq4-E226Q single mutants.



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 3.
Linear relationship between {Delta}A535 and NPQ. A, data from several individual leaves of each genotype. B, same data except that each point represents the mean ± S.E. for each genotype.

 


View this table:
[in this window]
[in a new window]
  		TABLE I
Statistical analysis of the results in Fig. 3

 
DCCD Binding—DCCD binds to carboxylate residues in hydrophobic environments (17, 37). It is known for its ability to inhibit qE (14–16), which suggests that carboxylate residues are involved in the process. PsbS purified from spinach or Arabidopsis and PsbS expressed in Escherichia coli bind DCCD at pH 7.5. Three acidic residues in each of the two lumen-exposed loops were suggested to be the possible binding sites (18). Fig. 4 shows that, in thylakoids isolated from wild-type Arabidopsis, DCCD bound to PsbS at pH 5. However, no binding of DCCD was detected at pH 7.8 (data not shown). DCCD binding was only 45 ± 10 and 50 ± 12% of the control in the npq4-E122Q and npq4-E226Q single mutants, respectively, and binding was undetectable in the npq4-E122Q,E226Q double mutant (Fig. 4). These results indicate that DCCD can equally bind to Glu-122 and Glu-226 and that mutation of one of the glutamates does not affect the chemical characteristics of the other in a cooperative manner.



View larger version (74K):
[in this window]
[in a new window]
  		FIG. 4.
[14C]DCCD binding to PsbS at pH 5.0. After separating thylakoid proteins by two-dimensional PAGE, the gel was stained with Coomassie Blue and then dried, and the radioactivity was detected with the Instant Imager (Packard). Thylakoids from plants carrying different mutations affecting PsbS were used for the labeling. The circles on the gel panel indicate other unidentified proteins labeled by [14C]DCCD in all thylakoids.

 
Chl a Fluorescence Lifetimes—We further analyzed the Chl a fluorescence lifetime distributions in these mutants. Fig. 5A shows the phase shift and demodulation ratio data as a function of modulation frequency (symbols) and the globally fit model (lines), with the residual errors in the lower panel. The patterns of the phase angle shifts and higher demodulation ratios indicate that the npq4–1 + psbS overexpression lines have the fastest decay times (lowest phase angles and highest demodulation ratios), followed by the wild type, the two single mutants npq4-E122Q and npq4-E226Q, the double mutant, and finally the npq4-1. The residual error plot indicates the model exhibits no remarkable systematic deviations and a random distribution around the mean = 0 for both the phase angle shifts and demodulation ratios.



View larger version (24K):
[in this window]
[in a new window]
  		FIG. 5.
Multifrequency phase and modulation time-resolved Chl a fluorescence data and simulated model interpretation from leaves of wild-type Arabidopsis and PsbS mutants. A, phase (angle) shifts and demodulation ratios as a function of modulation frequency for each type of leaf under conditions of saturating lumen acidity and subsaturating PSII photochemistry. The lower panel plots the residual errors for the phase shift (phase res) and demodulation ratios (demod res). B, lifetime-weighted fractional intensity distribution (f{tau}({tau})) as a function of the fluorescence lifetime, {tau}. The five positive (c1 and c3–c6) and one negative (c2) amplitude modes in the global model solution are indicated. Integration of the f{tau}({tau}) distribution for each sample yields the average lifetime <{tau}>.

 
Fig. 5B and Table II, respectively, show and tabulate the lifetime-weighted fractional intensity distributions as a function of the fluorescence lifetime and their integral contributions to the average lifetime for the five positive amplitude components (c1 and c3–c6) and one negative amplitude component (c2) used in the global analysis. There were two main positive fraction components, namely c6 and c4, in the npq4-1 and npq4-E122Q,E226Q double mutants that both lacked PsbS function. There were two positive components (c3 and c5) that correlated with PsbS function in all the other lines. The negative component c2 (centered ~140 ps) was weakest in the npq4-1 mutant that lacked PsbS and in the wild type that has only 20% of the PsbS protein level of the other mutant lines (Fig. 1). The other lines all exhibited larger c2 components. This observation suggests the negative c2 component was possibly more influenced by the PsbS protein level as opposed to the PsbS functional activity. The most rapid c1 component (<20 ps) was present in all PsbS-protein containing lines, and lowest in npq4-1. Consistent with previous results (33), c1 was stronger in the npq4-1 + psbS lines than in the wild type but appeared to be independent of the PsbS function, because it was similarly resolved in the single and double mutants, which have roughly the same levels of PsbS protein as the npq4–1 + psbS.


View this table:
[in this window]
[in a new window]
  		TABLE II
Fluorescence lifetime distribution component parameters

Data represent the integral lifetime-weighted fractional intensity contributions of each component in units of ps as the mean of three measurements with separate leaf samples. Confidence intervals were defined by S.D. of less than 10% of the average lifetime values for each mutant. Component distributions were as identified in Fig. 5 under steady-state PSII fluorescence conditions as defined previously (34).

 
With respect to the effect of PsbS on PSII photochemical efficiency, the npq4-1 and npq4-E122Q,E226Q double mutant lifetime distributions were predominated by c6 with the longest, broadest lifetime (~1100 ps), attributed to fluorescence from closed PSII traps (Fm), lesser contributions of the c4 component (~300 ps) that are likely mainly associated with open PSII trap (Fo) emission. The npq4-1 showed significantly higher amplitudes of c6 (and less c4) than the double mutant. The larger c4 component in the double mutant compared with the npq4-1 coincided with a larger negative c2 component. The two single mutants both exhibited nearly identical patterns for the distribution components with levels of c6 roughly 50% of those exhibited in the double mutant and also reduced levels of c4. The two single mutants also exhibited the two narrow components, c5 centered at ~525 ps and c3 at ~220 ps, which were clearly dependent on PsbS function and most likely originated from closed and open PSII traps, respectively. The wild type exhibited about 10% of the c6 level exhibited in the npq4-1 and was predominated by the c5 component. The npq4-1 + psbS was, similar to the wild type + psbS lines reported earlier (33), dominated by the c1, c3, and c5 components indicating the strongest energy dissipation and highest degree of PSII trap opening (largest c3 contribution), leading to the fastest average decay time in all lines compared in Fig. 5A. The npq4-1 + psbS exhibited less than 3% of the c6 level in the double mutant, indicating the PsbS functionality may have been nearly saturated. It was clear, however, that the npq4-1 + psbS line had slightly less PsbS activity than the wild type + psbS lines described previously (33). The wild type + psbS lines with higher PsbS protein levels exhibited larger relative c1:c3 ratios than the npq4-1 + psbS line analyzed here, leading to about a 56 ps (16%) faster average lifetime in the former samples.

Fig. 6 compares the intensity ratios measured with a PAM Chl fluorimeter during and after (Fmr/F) maximal levels of light-induced fluorescence quenching, along with the calculated average lifetime <{tau}> values under the Fs conditions from the experiments described in the legend to Fig. 5. The <{tau}> at Fs correlated strongly with the values in all lines. The npq4-1 and double mutant exhibited close parallels, with both the largest <{tau}> values (1.132 ns and 0.957 ns) and F' m/Fm values (0.54 and 0.52). The single mutants exhibited slightly higher <{tau}> and values (0.665 ns and 0.39) than the wild type (0.577 ns and 0.34) and the npq4-1 + psbS line showed the lowest values (0.386 ns and 0.23). Both the npq4-1 and the double mutant exhibited ~60% recovery of the original Fm, the two single mutants each exhibited about 65%, whereas the wild type and npq4-1 + psbS showed ~70 and 73%, respectively.



View larger version (34K):
[in this window]
[in a new window]
  		FIG. 6.
Comparison of steady-state PSII Chl a fluorescence ratios during and average fluorescence lifetimes from leaves of wild-type Arabidopsis and PsbS mutants. represents the steady-state Chl a fluorescence ratio under conditions of qE. <{tau}> is the average lifetime from fluorescence lifetime measurements. Fmr/Fm represents the steady-state Chl a fluorescence ratio after relaxation of qE.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
PsbS Senses the {Delta}pH through Two Symmetrically Arranged, Lumen-exposed Glutamates, Glu-122 and Glu-226 —Unlike typical light-harvesting complex proteins, PsbS has four rather than three transmembrane helices. Protein sequence analysis showed high similarity between helix I and helix III and also between helix II and helix IV (38, 39), reflecting the symmetrical topology of the PsbS protein (Fig. 7). The two lumen-exposed loops (40) are also highly similar, with Glu-122 and Glu-226 located in the middle of each loop. Mutating one or the other of these two glutamates only partly inhibits the PsbS function in qE, whereas the npq4-E122Q,E226Q double mutant totally disrupts the PsbS function (Fig. 2). These results are consistent with the suggestion that glutamates Glu-122 and Glu-226 of PsbS serve as the pH sensors for qE (13).



View larger version (31K):
[in this window]
[in a new window]
  		FIG. 7.
Topology of PsbS. Symmetrically arranged amino acid residues in the first half (dark shading) and second half (gray shading) of the protein are highlighted. The two glutamates discussed in this paper are denoted by square symbols and numbered.

 
The function of Glu-122 and Glu-226 was further confirmed by investigation of DCCD binding. DCCD is a protein modifying reagent that covalently binds to carboxylate residues involved in reversible protonation in hydrophobic environments (17). DCCD has previously been shown to act as a specific inhibitor of qE (14–16), thus leading to the search for target sites using [14C]DCCD. Binding sites have been identified in Lhcb proteins (41), namely CP26 (42) and CP29 (43), but evidence for these proteins being the functional binding site(s) of this qE inhibitor was lacking. Our results show that the single mutations E122Q and E226Q each reduce the level of DCCD binding to PsbS by ~50% and in the same way decrease the amplitude of qE. These findings imply that PsbS, rather than CP26 or CP29, is the target site for qE inhibition by DCCD. Moreover, because the double mutant is not labeled, Glu-122 and Glu-226 are the only DCCD binding sites in PsbS, implying that they have an essential role in PsbS function. It should be noted that purified PsbS, either extracted from thylakoids or recombinant from E. coli, binds DCCD at pH 7.5, but low pH is required for labeling of PsbS in thylakoid membranes, suggesting that either the conformation of PsbS is different in the two environments or that interactions with neighbor subunits in PSII supercomplexes prevents exposure of lumenal glutamate residues until the pH is decreased.

It is interesting to note that the two glutamate residues are responsible each for 50% of both DCCD binding and control of qE activity. This suggests that, whatever the mechanism is for the activity of PsbS in qE, the two halves of the 2-fold symmetrical molecule are acting independently. Recent work (12) showed that in the presence of PsbS, a 10-ps spectral component is present that can be interpreted as a Chl a to Z energy transfer, and Z binding to PsbS has also been reported (23). On this basis, the qE quenching can be tentatively understood in terms of the protonation of each glutamate leading to binding of two Z molecules, which are capable of accepting and dissipating energy from singlet excited Chl a. Thus, PsbS can be viewed as a thylakoid sensor of excess light. In the presence of a low thylakoid lumen pH, protonation of PsbS results in thermal dissipation of excess absorbed light energy, i.e. a feedback down-regulation of photosynthetic light harvesting by the {Delta}pH.

In addition to influencing qE, the E122Q and E226Q mutations also inhibit the {Delta}A535 leaf absorbance change (Fig. 3). It was found early on that {Delta}A535 is invariably associated with qE (20, 21). Here we report that a single linear relationship exists between NPQ and {Delta}A535 even when taking into consideration two different factors, namely different levels of PsbS protein and different amino acid substitutions in the PsbS protein (Fig. 3). It was hypothesized that {Delta}A535 is because of a red absorption shift of Z upon binding to PsbS (22). The linear relationship between {Delta}A535 and NPQ suggests that one of two Z-binding sites is affected in each single mutant (E122Q or E226Q), whereas Z binding might be eliminated completely in the double mutant (E122Q,E226Q).

Interpreting the Fluorescence Lifetime Distribution Components with Respect to the Glutamate Mutants, PsbS Protein Level, and PSII Function—As illustrated in Fig. 8A, our current biochemical model of PsbS function postulates that PsbS associated with PSII may exist in one of three states depending on the lumen acidity and the concentration of Z (and/or A). The W state is defined as a PsbS with its critical glutamate residues (Glu-122 and Glu-226) in an unprotonated state. W converts reversibly to X upon protonation of the glutamates as determined by their pKa value. The protonated state X is hypothesized to be an activated state for potential binding of Z. Upon binding of Z, the PsbS switches from the X to the Y state, which is the fully functional energy dissipation mode.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 8.
Biochemical scheme of the lumen pH- and Z-dependent effects on the PsbS protein with respect to energy dissipation and reaction center photochemistry in PSII. A, model scheme with H+ and Z binding steps (24). B, interpretation of the PSII fluorescence lifetime distribution components with respect to the combined influence of energy dissipation and reaction center photochemistry. Wo (Wc), Xo (Xc), and Yo (Yc) represent PSII in the W, X, and Y states (defined in panel A) with the redox state of the primary PSII electron acceptor QA in an oxidized (reduced) state. The lifetime components c3–c6 were as defined in Fig. 5.

 
Consistent with our recent papers (33, 44), we have found it necessary to expand our model to six states to account for the opening/closing (oxidation/reduction) of the PSII reaction centers. To simplify the model parameterization and interpretation, our experiments were designed to yield both saturating light-induced lumen pH and Z concentration conditions, thereby reducing to a minimum both the W (by saturated protonation) and X states (by saturated xanthophyll binding). Practically, we have also concluded that we are unable to resolve the W and X components in either the open or closed PSII states, most likely because they are likely to differ by less than 20% between the open and closed W or X components. Therefore, our empirical global model linking scheme assumes the Wc + Xc and Wo + Xo components to be indistinguishable, yielding a simplified four-component scheme (Fig. 8B). As mentioned in the "Results," we postulate the rapid c1 and negative c2 components to be more strongly related to the PsbS protein level than PsbS function per se, because these components were resolved in the double and both single mutants. Hence, we do not consider them to be associated directly with a particular state of PSII in the above scheme. We propose that extremely high PsbS concentrations may have indirect effects on the photochemical activity of the PSII core antenna and possibly alter energy transfer pathways among Chl complexes, perhaps between the antenna and core. With respect to Fig. 8B, the (1100 ps) c6 component of our lifetime distribution model would be assigned to the combined [Wc + Xc] states. We emphasize that the assignment of c6 to the W state is supported by the fact that the c6 contribution is maximal in the npq4-1, very similar to npq4-1 in the double mutant, reduced by ~50% in both single mutants (E122Q and E226Q) compared with the double mutant, reduced by >80% in the wild type and down to less than 3% in the npq4–1 + psbS. The c4 (300 ps) component appears consistent with emission from the Wo + Xo state PsbS conformations with open PSII traps. However, as indicated above it is unlikely that the Wo state would be a significant component because the key PsbS glutamate residues were likely titrated to virtually complete protonation. Likewise, we feel it is logical to assign the c3 (220 ps) and c5 (525 ps) states to the open Yo and closed Yc PSII centers, respectively. A main tenet of the model outlined in Fig. 8B that is consistent with many steady-state spectroscopy experiments, is that increasing the population of the Y state acts synergistically to yield a higher relative fraction of open/oxidized PSII traps (Yo) with enhanced levels of energy dissipation. The increased rate constant of dissipation of excess absorbed light in the open and closed PSII traps reduces the "excitation pressure" on the PSII reaction center Chls leading to a higher population of open PSII traps (10, 33, 44).

The c6 mode was ~600 ps shorter than observed in thylakoid experiments (W = 1740 ps). This observation is consistent with the larger decrease of the Fm (30–40%) following the illumination treatments in these leaf experiments (1250 µmol photons m–2 s–1 for 20 min) as opposed to 20–25% observed in isolated thylakoids that were treated with lower (500 µmol photons m–2 s–1) light intensities (36). We note as mentioned before that in thylakoids we measured an additional lumen pH-induced 16% decrease in the lifetime center of the W mode tentatively attributed to protonation and/or conformational changes of other PSII proteins besides PsbS, which was absent in the npq4-1 mutant. Previously, we reported that the W mode center was decreased during aerobic photoinhibition in thylakoids largely independent of the width or fraction of the distribution (45). So both things considered, we conclude that the ~50% decrease in the W mode in the condition compared with W in the Fm state in these leaf experiments is attributed to both photoinhibitory damage caused by the strong light and protonation/conformational changes in other PSII proteins besides PsbS. Furthermore, we consider that all lifetime modes may be partially attenuated by PSII photochemical activity through kinetic processes that are beyond the resolution of our instrument (i.e. 5–10 ps).

Concluding Remarks—One of the key points of the model described above is the implications of the lifetime distribution fractional intensities on the structure-function of PsbS. In our view the simplest explanation for the similarities of the two single mutants (E122Q and E226Q) is that they have equivalent effects because of the bilateral symmetry of the PsbS protein structure. The data indicate, as suggested in Fig. 9, that each single mutation elicits a ~50% inhibition effect that becomes 100% in the double mutant. Hence, we propose that PsbS has two equivalent functional sites, one on each side, which are likely associated with pH-activated binding of the xanthophylls. It is thus suggested that PsbS associates with its binding site to attach itself to the PSII holocomplex (likely in or near the core antenna proteins) in one of two facial orientations with a 50% probability. And furthermore, according to this model, two Z per PsbS will be the saturation amount for qE.



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 9.
Model explaining the effect of PsbS glutamate mutants on H+ and Z binding and proposed interactions between PsbS and PSII. The glutamate to glutamine mutations of PsbS are represented by the -CONH2 moieties, and their lack of function in qE is represented by the black ovals (putative xanthophyll binding sites) to which they are attached. The -COO– and -COOH moieties represent the charged and protonated glutamates, respectively. Upon protonation, a xanthophyll binding site is generated, as indicated by the change from the gray (-COO–) to the white ovals (-COOH). The presence of a xanthophyll molecule (Z, A, or possibly lutein) occupying a protonated binding site is shown in the Y state. In the double mutant npq4-E112Q,E226Q shown here and in npq4-1 (not shown), protonation of PsbS does not occur, and it is assumed that [W] = 1. In the single mutants npq4-E122Q or npq4-E226Q, two possible interactions of symmetrical PsbS with PSII are shown, one of which is a functional interaction. In this case, [W] would be 0.5. For npq4-1 + psbS (and in the wild type, not shown), either possible interaction orientation of PsbS with PSII is functional.

 

   FOOTNOTES
 
* This work was supported by the Director, Office of Science, Office of Basic Energy Sciences, Chemical Sciences Division, of the United States Department of Energy under contract number DE-AC03-76SF00098 (to K. K. N.), and Grants A00103734 and DP0343160 from the Australian Research Council (to A. M. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Both authors contributed equally to this paper. Back

|| Present address: Jobin-Yvon Inc., 3880 Park Ave., Edison, NJ 08820. Back

§§ Supported by European Community Human Potential Program contract HPRN-CT-2002-00248 (PSICO). Back

|||| To whom correspondence should be addressed. Tel.: 510-643-6602; Fax: 510-642-4995; E-mail: niyogi{at}nature.berkeley.edu.

1 The abbreviations used are: qE, rapidly inducible, pH- and xanthophyll-dependent component of NPQ; A, antheraxanthin; Chl, chlorophyll; DCCD, N,N'-dicyclohexylcarbodiimide; NPQ, nonphotochemical quenching of chlorophyll fluorescence; PSII, photosystem II; V, violaxanthin; Z, zeaxanthin; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine. Back

2 Available at math.nist.gov. Back


   ACKNOWLEDGMENTS
 
We thank Anastasios Melis for providing the anti-D1 antibody.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Horton, P., Ruban, A. V., and Walters, R. G. (1996) Annu. Rev. Plant Physiol. Plant Mol. Biol. 47, 655–684[CrossRef]
  2. Gilmore, A. M. (1997) Physiol. Plant. 99, 197–209[CrossRef]
  3. Niyogi, K. K. (1999) Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 333–359[CrossRef]
  4. Müller, P., Li, X.-P., and Niyogi, K. K. (2001) Plant Physiol. 125, 1558–1566[Free Full Text]
  5. Gilmore, A. M., Hazlett, T. L., and Govindjee (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 2273–2277[Abstract/Free Full Text]
  6. Yamamoto, H. Y., Nakayama, T. O., and Chichester, C. O. (1962) Arch. Biochem. Biophys. 97, 168–173
  7. Demmig-Adams, B. (1990) Biochim. Biophys. Acta 1020, 1–24[CrossRef]
  8. Gilmore, A. M., and Yamamoto, H. Y. (1993) Photosynth. Res. 35, 67–78[CrossRef]
  9. Niyogi, K. K., Grossman, A. R., and Björkman, O. (1998) Plant Cell 10, 1121–1134[Abstract/Free Full Text]
  10. 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]
  11. Frank, H. A., Cua, A., Chynwat, V., Young, A., Gosztola, D., and Wasielewski, M. R. (1994) Photosynth. Res. 41, 389–395[CrossRef]
  12. Ma, Y. Z., Holt, N. E., Li, X.-P., Niyogi, K. K., and Fleming, G. R. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 4377–4382[Abstract/Free Full Text]
  13. Li, X.-P., Phippard, A., Pasari, J., and Niyogi, K. K. (2002) Funct. Plant Biol. 29, 1131–1139[CrossRef]
  14. Ruban, A. V., Walters, R. G., and Horton, P. (1992) FEBS Lett. 309, 175–179[CrossRef][Medline] [Order article via Infotrieve]
  15. Crofts, A. R., and Yerkes, C. T. (1994) FEBS Lett. 352, 265–270[CrossRef][Medline] [Order article via Infotrieve]
  16. Walters, R. G., Ruban, A. V., and Horton, P. (1994) Eur. J. Biochem. 226, 1063–1069[Medline] [Order article via Infotrieve]
  17. Azzi, A., Casey, R. P., and Nalecz, M. J. (1984) Biochim. Biophys. Acta 768, 209–226[Medline] [Order article via Infotrieve]
  18. Dominici, P., Caffarri, S., Armenante, F., Ceoldo, S., Crimi, M., and Bassi, R. (2002) J. Biol. Chem. 277, 22750–22758[Abstract/Free Full Text]
  19. Heber, U. (1969) Biochim. Biophys. Acta 180, 302–319[Medline] [Order article via Infotrieve]
  20. Bilger, W., Björkman, O., and Thayer, S. S. (1989) Plant Physiol. 91, 542–551[Abstract/Free Full Text]
  21. Bilger, W., and Björkman, O. (1994) Planta 193, 238–246
  22. Ruban, A. V., Pascal, A. A., Robert, B., and Horton, P. (2002) J. Biol. Chem. 277, 7785–7789[Abstract/Free Full Text]
  23. Aspinall-O'Dea, M., Wentworth, M., Pascal, A., Robert, B., Ruban, A., and Horton, P. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 16331–16335[Abstract/Free Full Text]
  24. Gilmore, A. M., Shinkarev, V. P., Hazlett, T. L., and Govindjee (1998) Biochemistry 37, 13582–13593[CrossRef][Medline] [Order article via Infotrieve]
  25. Müller-Moulé, P., Conklin, P. L., and Niyogi, K. K. (2002) Plant Physiol. 128, 970–977[Abstract/Free Full Text]
  26. Sacksteder, C. A., Jacoby, M. E., and Kramer, D. M. (2001) Photosynth. Res. 70, 231–240[CrossRef][Medline] [Order article via Infotrieve]
  27. Avenson, T., Cruz, J. A., and Kramer, D. M. (2004) Proc. Natl. Acad. Sci. U. S. A. 101, 5530–5535[Abstract/Free Full Text]
  28. Sacksteder, C. A., Kanazawa, A., Jacoby, M. E., and Kramer, D. M. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 14283–14288[Abstract/Free Full Text]
  29. Bassi, R., Rigoni, F., Barbato, R., and Giacometti, G. M. (1988) Biochim. Biophys. Acta 936, 29–38[CrossRef]
  30. Bassi, R., Machold, O., and Simpson, D. (1985) Carlsberg Res. Commun. 50, 145–162
  31. Laemmli, U. K. (1970) Nature 227, 680–685[CrossRef][Medline] [Order article via Infotrieve]
  32. Gilmore, A. M., and Björkman, O. (1995) Planta 197, 646–654
  33. Li, X.-P., Müller-Moulé, P., Gilmore, A. M., and Niyogi, K. K. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 15222–15227[Abstract/Free Full Text]
  34. van Kooten, O., and Snel, J. F. H. (1990) Photosynth. Res. 25, 147–150[CrossRef]
  35. Gilmore, A. M. (2004) in Chlorophyll Fluorescence: A Signature of Photosynthesis (Papageorgiou, G. C., and Govindjee, eds) Kluwer Academic Publishers, Dordrecht, The Netherlands, in press
  36. Li, X.-P., Gilmore, A. M., and Niyogi, K. K. (2002) J. Biol. Chem. 277, 33590–33597[Abstract/Free Full Text]
  37. Hoppe, J., and Sebald, W. (1980) Eur. J. Biochem. 107, 57–65[Medline] [Order article via Infotrieve]
  38. Green, B. R., and Pichersky, E. (1994) Photosynth. Res. 39, 149–162[CrossRef]
  39. Jansson, S. (1999) Trends Plant Sci. 4, 236–240[CrossRef][Medline] [Order article via Infotrieve]
  40. Kim, S., Pichersky, E., and Yocum, C. F. (1994) Biochim. Biophys. Acta 1188, 339–348[Medline] [Order article via Infotrieve]
  41. Jahns, P., and Junge, W. (1990) Eur. J. Biochem. 193, 731–736[Medline] [Order article via Infotrieve]
  42. Walters, R. G., Ruban, A. V., and Horton, P. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 14204–14209[Abstract/Free Full Text]
  43. Pesaresi, P., Sandona, D., Giuffra, E., and Bassi, R. (1997) FEBS Lett. 402, 151–156[CrossRef][Medline] [Order article via Infotrieve]
  44. Gilmore, A. M. (2001) Photosynth. Res. 67, 89–101
  45. Gilmore, A. M., Hazlett, T. L., Debrunner, P. G., and Govindjee (1996) Photochem. Photobiol. 64, 552–563[Medline] [Order article via Infotrieve]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 ScienceHome page
T. K. Ahn, T. J. Avenson, M. Ballottari, Y.-C. Cheng, K. K. Niyogi, R. Bassi, and G. R. Fleming
Architecture of a Charge-Transfer State Regulating Light Harvesting in a Plant Antenna Protein
Science, May 9, 2008; 320(5877): 794 - 797.
[Abstract] [Full Text] [PDF]

Home page
 Plant CellHome page
S. de Bianchi, L. Dall'Osto, G. Tognon, T. Morosinotto, and R. Bassi
Minor Antenna Proteins CP24 and CP26 Affect the Interactions between Photosystem II Subunits and the Electron Transport Rate in Grana Membranes of Arabidopsis
PLANT CELL, April 1, 2008; 20(4): 1012 - 1028.
[Abstract] [Full Text] [PDF]

Home page