Regulating the proton budget of higher plant photosynthesis.
ABSTRACT In higher plant chloroplasts, transthylakoid proton motive force serves both to drive the synthesis of ATP and to regulate light capture by the photosynthetic antenna to prevent photodamage. In vivo probes of the proton circuit in wild-type and a mutant strain of Arabidopsis thaliana show that regulation of light capture is modulated primarily by altering the resistance of proton efflux from the thylakoid lumen, whereas modulation of proton influx through cyclic electron flow around photosystem I is suggested to play a role in regulating the ATP/NADPH output ratio of the light reactions.
- SourceAvailable from: pnas.orgProceedings of the National Academy of Sciences 11/2002; 99(20):12518-9. · 9.74 Impact Factor
- [show abstract] [hide abstract]
ABSTRACT: The F(1)F(0)-type ATP synthase is a key enzyme in cellular energy interconversion. During ATP synthesis, this large protein complex uses a proton gradient and the associated membrane potential to synthesize ATP. It can also reverse and hydrolyze ATP to generate a proton gradient. The structure of this enzyme in different functional forms is now being rapidly elucidated. The emerging consensus is that the enzyme is constructed as two rotary motors, one in the F(1) part that links catalytic site events with movements of an internal rotor, and the other in the F(0) part, linking proton translocation to movements of this F(0) rotor. Although both motors can work separately, they must be connected together to interconvert energy. Evidence for the function of the rotary motor, from structural, genetic and biophysical studies, is reviewed here, and some uncertainties and remaining mysteries of the enzyme mechanism are also discussed.Trends in Biochemical Sciences 04/2002; 27(3):154-60. · 13.08 Impact Factor
- Biochimica et Biophysica Acta 04/1979; 505(3-4):355-427. · 4.66 Impact Factor
Regulating the proton budget of higher
Thomas J. Avenson, Jeffrey A. Cruz, Atsuko Kanazawa, and David M. Kramer*
Institute of Biological Chemistry, 289 Clark Hall, Washington State University, Pullman, WA 99164-6340
Communicated by Clarence A. Ryan, Washington State University, Pullman, WA, May 12, 2005 (received for review February 7, 2005)
In higher plant chloroplasts, transthylakoid proton motive force
serves both to drive the synthesis of ATP and to regulate light
capture by the photosynthetic antenna to prevent photodamage.
of Arabidopsis thaliana show that regulation of light capture is
modulated primarily by altering the resistance of proton efflux
from the thylakoid lumen, whereas modulation of proton influx
through cyclic electron flow around photosystem I is suggested to
play a role in regulating the ATP?NADPH output ratio of the light
ATP synthase proton conductivity ? cyclic electron flow ? linear electron
flow ? energy-dependent nonphotochemical quenching ?
protein motive force
Higher plant photosynthesis is initiated through absorption of
light by antennae complexes that funnel the energy to photo-
system (PS) II and I. The photosystems operate in sequence with
the plastoquinone pool, the cytochrome b6f complex, and plas-
tocyanin to oxidize H2O and reduce NADP?to NADPH in what
is termed linear electron flow (LEF). LEF is coupled to proton
translocation, establishing a transthylakoid electrochemical gra-
dient of protons, termed the proton motive force (pmf) (2),
comprised of electric field (??) and pH (?pH) gradients (3).
hotosynthesis converts light energy into chemical energy,
ultimately powering the vast majority of our ecosystem (1).
Dual Role of the pmf
The pmf plays two central roles in higher plant photosynthesis
(4). First, pmf drives the normally endergonic synthesis of ATP
through the CF1-CF0ATP synthase (ATP synthase) (5). Both
the ?pH and ?? components of pmf contribute to ATP synthesis
in a thermodynamically, and probably kinetically, equivalent
fashion (6). Second, pmf is a key signal for initiating photopro-
tection of the photosynthetic reaction centers through energy-
dependent nonphotochemical quenching (qE), a process that
harmlessly dissipates excessively absorbed light energy as heat
(7–10). Only the ?pH component of pmf, through acidification
of the lumen, is effective in initiating qEby activating violaxan-
thin de-epoxidase, a lumen-localized enzyme that converts vio-
laxanthin to antheraxanthin and zeaxanthin, and by protonating
lumen-exposed residues of PsbS, a pigment-binding protein of
the PS II antenna complex (11).
A Need for Flexibility in the Light Reactions
A major open question concerns how the light reactions achieve
the flexibility required to meet regulatory needs and match
downstream biochemical demands (12). In LEF to NADP?, the
synthesis of ATP and the production of NADPH are coupled,
producing a fixed ATP?NADPH output ratio. LEF alone is
probably unable to satisfy the variable ATP?NADPH output
ratios required to power the sum of the Calvin–Benson cycle (13,
14) and other metabolic processes (alternate electron and ATP
sinks) that are variably engaged under different physiological
conditions (12, 15, 16). Failure to match ATP?NADPH output
with demand will lead to buildup of products and depletion of
substrates for the light reactions, leading to inhibition of the
The generation of pmf is likewise coupled to LEF, so it is clear
that the sensitivity of antenna regulation (or qE) must also be
modulated in some way to avoid catastrophic failure of photo-
protection (12, 15, 17–19). Longer-term acclimation of the qE
response can involve altering the sensitivity of the regulatory
machinery to lumen pH by changing the xanthophyll pigment
and?or PsbS levels (12, 20). However, dramatic changes in light
intensity and?or CO2availability can occur over the seconds-
to-hours time scale (8), requiring short-term adjustments. In-
or O2levels can strongly modulate (by up to 6-fold) the sensi-
tivity of qEwith respect to LEF (17, 18).
Two Types of Flexibility Mechanisms
Two general types of models have been proposed to account for
the flexibility required to meet these changing demands (12). In
‘‘Type I’’ mechanisms, proton flux into the lumen is increased
through alternate electron transfer pathways, especially cyclic
electron flow associated with PSI (CEF1), a mechanism that
returns electrons from PSI to the plastoquinone pool, thereby
increasing the magnitude of the pmf relative to that generated by
LEF alone (12). Other processes are also possible, for example,
turnover of a plastid terminal oxidase (21, 22), but these
processes would have to run at relatively high rates to signifi-
cantly impact the overall ATP?NADPH balance. For C3vascular
plants, CEF1 has been suggested to supply the relatively small
fluxes (10–15% of that supplied by LEF) of protons required to
balance ATP?NADPH output for the Calvin–Benson cycle and
24) as to whether CEF1 can run at sufficiently high rates to alter
qEresponses by up to 6-fold, especially given the expected large
ATP?NADPH imbalances such large fluxes would likely incur
In Type II mechanisms, lumen acidification with respect to
the lumen, thus modulating qE sensitivity without impacting
ATP?NADPH output. This phenomenon is thought to be
achieved by varying either the conductivity of the CF1-CF0 ATP
synthase to proton efflux as measured by electrochromic shift
(ECS) decay (gH?), i.e., the inverse of the resistance to proton
efflux from the lumen or the relative fraction of pmf stored as
?pH (12, 16–18, 24).
Probing the pmf to Gain Insight into the
Recently, a series of in vivo probes of the pmf have been
introduced (2, 3, 16, 25–28), allowing contributions from Types
I and II flexibility mechanisms to be directly assessed. These
Abbreviations: CEF1, cyclic electron flow associated with PSI; ECS, electrochromic shift; LC,
low CO2(50 ppm CO2, 21% O2); LEF, linear electron flow; PS, photosystem; pmf, proton
motive force; pmfLEF, pmf generated by LEF; qE, energy-dependent nonphotochemical
*To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
© 2005 by The National Academy of Sciences of the USA
July 5, 2005 ?
vol. 102 ?
no. 27 ?
techniques are based on kinetic analyses of the ECS (26) of
photosynthetic pigments, which yields absorbance changes pro-
portional to changes in transthylakoid ?? (29). Several useful
parameters can be obtained from analysis of ECS decay kinetics
during brief dark perturbations of the steady state, including
estimates of the relative flux of protons through the ATP
synthase (?H?, which at steady state equals flux of protons into
the lumen), the magnitude of the light-induced pmf, the fraction
of pmf stored as ?pH and ??, and gH?(3, 16–18, 25, 26, 28).
Combined with standard chlorophyll a fluorescence assays, from
which estimates of LEF can be obtained (30), one can calculate
the pmf generated by LEF alone (i.e., pmfLEF? LEF?gH?), a key
parameter for estimating fractional changes in CEF1 turnover
Using these probes of the proton circuit, it was shown that in
intact Nicotiana tabacum (tobacco) leaves, lowering atmospheric
CO2 from 372 to 0 ppm led to a ?5-fold increase in the
dependence of qE on LEF (17). The effect could be entirely
accounted for by a proportional (i.e., 5-fold) decrease in gH?, so
that even modest rates of LEF generated a substantial pmf and
a robust qEresponse (17, 18). A similar (?6-fold) change in qE
sensitivity was observed when both O2and CO2were lowered (to
1% and 50 ppm, respectively), but in this case, both changes in
gH?and increased partitioning of pmf into ?pH were invoked to
explain the effect (18). In both cases, the ratio of vH??LEF
remained essentially constant (within noise levels), indicating
that contributions from CEF1 to proton flux were either small
or remained a relatively constant fraction of those from LEF, as
previously found for tobacco (25). On the whole, these results
support a large role for Type II mechanisms in modulating qE
sensitivity upon short term changes in CO2?O2levels, but they
do not rule out smaller contributions from Type I mechanisms
in balancing ATP?NADPH output (12, 16, 28).
partial characterization of a mutant strain of Arabidopsis thali-
ana, termed pgr5 for proton gradient regulation, which showed
two provocative phenotypes. First, nonphotochemical reduction
of the plastoquinone pool, attributed to the key step in CEF1,
was inhibited in pgr5. Second, qEwas severely diminished. It is
reasonable to hypothesize that the loss of PGR5 blocks CEF1
and, thereby, abolishes a significant flux of protons needed to
activate qE (31, 32). Evidence for such a hypothesis would
support a large role for Type I mechanisms in modulating qE
sensitivity (33) while arguing against Type II models (12, 17, 18).
On the other hand, mutation of pgr5 could indirectly affect qEby
disrupting downstream processes and modulating metabolic
pool sizes (31, 32). Here, we present an experimental test for
causal links between the loss of PGR5, steady-state proton flux,
and the qEresponse, allowing us to determine the relative roles
of Type I and II flexibility responses.
Materials and Methods
Plant Strains and Growth Conditions. Wild-type A. thaliana (Wt-
background strain gl1) (31) and pgr5 plants were grown in tightly
controlled chambers under a 16:8 photoperiod at an average of
?70 ?mol photons m?2s?1photosynthetically active radiation
and at 23°C. These growth conditions stably reproduced the
phenotypes seen previously over the entire experimental period
(31). Wt (gl1) and pgr5 seeds were a gift from T. Shikanai (Nara
Institute of Science and Technology, Ikoma, Nara, Japan).
Spectroscopic Assays. Fully expanded leaves from ?23- to 26-day-
old plants were used in spectroscopic assays. Room air (372 ppm
CO2?21% O2) or premixed gases from cylinders (i.e., 50 ppm
CO2?21% O2) were bubbled through water (for humidification)
before entering the measuring chamber of the spectrophotom-
eter. Leaves were clamped into the measuring chamber of a
specifically designed for use on leaves (17, 18, 34). Leaves were
first exposed to 26–216 ?mol photons m?2s?1photosynthetically
active radiation from a series of red light emitting diodes
(maximum emission wavelength of 637 nm) to reach steady-state
effect. After this actinic period, the steady-state (Fs) and light
saturated (FM?) levels of chlorophyll a fluorescence yield were
obtained (17, 18), from which estimates of the efficiency of PSII
photochemistry (?II) were calculated (30). Estimates of LEF
were obtained by using ?IIas in ref. 35. Analyses of the ECS
decay kinetics upon perturbation of the steady state with an
and 26. Previous assays showed linear correlations between
estimates of LEF taken from fluorescence and absorbance
measurements with our instruments (25–27, 34), suggesting that
the spectroscopic techniques probed similarly responding pop-
nm were recorded in series, and those attributable to changes in
ECS were deconvoluted from background signals according to
the following equation (25, 26):
?ECS ? ??I?I0?520?? ????I?I0?535?? ??I?I0?505???2?).
taken as the total amplitude of ECS decay from its steady-state
(16–18). Relative estimates of the conductivity of the thylakoid
membrane to protons (gH?), primarily attributable to the turn-
over of the ATP synthase, were obtained by taking the inverse
of the time constant for ECS decay (?ECS) (16–18, 28). Relative
estimates of the pmf attributable to proton flux from LEF,
termed pmfLEF, were calculated by using the following equation
(16, 18, 28):
This parameter estimates the light-driven proton flux through
the ATP synthase based on the extent of LEF and the kinetic
properties of the ATP synthase turnover, as reviewed in ref. 28.
Western Blot Analyses. Crude leaf extracts from Wt and pgr5 were
prepared as described in ref. 36. Flash-frozen tissue was ground
in a mortar and pestle before resuspension in SDS?PAGE
sample buffer. Ten micrograms of protein, as estimated by using
the BCA Protein Assay Kit (Pierce), from each preparation was
loaded onto an SDS?PAGE gel. Protein was transferred to
poly(vinyl difluoride) membranes and probed with antibody
directed against the ?-subunit of the ATP synthase (a gift from
Alice Barkan, University of Oregon, Eugene, OR). Immunore-
active bands were detected on radiographic film by using the
SuperSignal West Pico Chemiluminescent Substrate kit (Pierce).
Similar conclusions were reached when the gel was loaded with
10, 30, or 90 ?g of protein, indicating that the assay was within
the linear range of detection (data not shown).
Results and Discussion
Effects of Lowering CO2 Levels and Loss of PGR5 on LEF and qE
Sensitivity. Fig. 1A shows plots of qEas a function of LEF from
26–216 ?mol photons m?2s?1for the wild type (Wt, gl1) (31)
under ambient air (372 ppm CO2?21% O2) and two different
treatments that lowered light saturated LEF by about the same
extent. Low CO2 air (LC; 50 ppm CO2?21% O2) reduced
light-saturated LEF in Wt by ?30%, a typical response for
A. thaliana (35). A similar lowering of light-saturated LEF was
obtained by using pgr5 under ambient air. These conditions were
chosen to avoid significant photoinhibition, which appeared in
pgr5 at ?216 ?mol photons m?2s?1as a decrease in LEF, as well
www.pnas.org?cgi?doi?10.1073?pnas.0503952102Avenson et al.
as large changes in the partitioning of the pmf into ?? and ?pH,
a phenomenon that has been previously observed in N. tabacum
under severe stress (18). Under more extreme conditions (higher
light intensities or lower CO2levels), results were qualitatively
consistent with those presented here (data not shown) as long as
partitioning of pmf into ?? and ?pH was considered (18).
In Wt under ambient air, a flux of ?40 ?mol electrons m?2s?1
generated a qEof 0.4, whereas the same level of qEwas achieved
at a flux of ?27 ?mol electrons m?2s?1under LC air (Fig. 1A).
At saturating light, qEwas ?35% larger under LC than ambient
air, despite having a slower LEF. Thus, similar to previous
observations in N. tabacum (17, 18), lowering CO2 in Wt
increased the sensitivity of qEwith respect to LEF. In contrast,
the ?30% decrease in LEF that occurred in the absence of
PGR5 was not accompanied by a corresponding increase in the
light saturated qEresponse, but was rather 4- to 6-fold lower in
comparison with that in the Wt.
Effects of Lowering CO2Levels and Loss of PGR5 on Contributions of
CEF1 to the Proton Budget. In Wt, varying the CO2levels had no
observable effects on the relationship between ?H? and LEF
(Fig. 1B), arguing against large CO2-dependent changes in
contributions from Type I modulation (12, 16–18). On the other
hand, the slope of ?H?vs. LEF was ?13% smaller (P ? 0.05) in
pgr5 than in Wt (Fig. 1B). The results were not significantly
altered by forcing the linear fits through the origin. Although
these small differences could be the result of small systematic
errors, e.g., in LEF measurements (37), they are also consistent
with results from Munekage et al. (31, 32) that PGR5 is
important for steady-state CEF1, and we thus adopt this view as
our working model.
This model is supported in separate estimates of proton flux
and pmf. The data in Fig. 2 shows the relationships between
estimates of the pmf attributable solely to proton translocation
by LEF (pmfLEF) and the total pmf (ECSt), driven by the sum of
LEF and other process (i.e., CEF1). Within the noise level, the
relationships for Wt under the two CO2 levels overlapped
(analysis of covariance indicated no significant differences in
slopes, P ? 0.6), implying that either LEF accounted for the vast
majority of estimated pmf, or that contributions from other
processes (see above), most notably CEF1, were a constant
fraction of LEF. Again, the slope of pmfLEFversus ECStwas
?14% smaller in pgr5 in comparison to Wt under ambient
conditions, a difference that was statistically significant (analysis
of covariance, P ? 0.05).
It is important to note that the ECStestimate of pmf is based
on the light-dark difference in the amplitude of the ECS signal
(17, 18), whereas the pmfLEFestimate of pmf is based on ECS
decay kinetics (18), i.e., the latter is not sensitive to changes in
the absolute ECS response. The leaf contents of photosynthetic
complexes were equivalent in Wt and pgr5 (31), and the ampli-
tudes of the rapid (?1 ms) ECS responses after saturating, single
turnover flashes, which reflect charge separation in PSII and PSI
centers (38), were indistinguishable, with Wt and pgr5 giving
3.5 ? 0.35 and 3.5 ? 0.24 (?I?I0? 1,000) respectively, indicating
essentially identical responses to ??. Overall, the constancy of
across the thylakoid membrane. Chlorophyll a fluorescence yield and ECS
ing (qE) (A) and steady-state proton flux into the lumen (?H?) (B), respectively,
from 26 to 216 ?mol photons m?2s?1on leaves from A. thaliana Wt under
LEF (18). Linear regressions of LEF versus ?H? are shown in B, the regression
slopes of which are 2.035 (solid line), 2.038 (dotted line), and 1.774 (dashed
line) for Wt ambient air, Wt?LC air, and pgr5 ambient air, respectively. Slopes
for Wt?atmospheric and pgr5?atmospheric were judged by analysis of covari-
by LEF alone. ECS and chlorophyll a fluorescence yield analyses were per-
formed on leaves from A. thaliana Wt plants and pgr5 to estimate light-
induced pmf (ECSt) and LEF, respectively, from which estimates of the pmf
regressions of pmfLEFversus ECStare shown, the slopes of which are 1.972
(solid line), 2.053 (dotted line), and 1.701 (dashed line) for Wt?ambient air,
pgr5?atmospheric were ?14% different and judged by analysis of covariance
slopes of Wt?atmospheric versus Wt?LC was not statistically significant (P ?
The relationship between light-induced pmf and the pmf generated
Avenson et al.
July 5, 2005 ?
vol. 102 ?
no. 27 ?
these results supports the validity of comparisons of the ECS-
derived parameters between the two strains.
Differences in qE Sensitivity Between Wt and pgr5 Can Be Largely
Attributed to Changes in gH?. The above flux estimates suggest
differences in contributions to light-induced pmf from processes
other than LEF, consistent with a difference in CEF1 engage-
ment between Wt and pgr5 (31, 32). However, the modest
(?13%) decrease in ?H? in the absence of PGR5 was far too
small to directly account for the corresponding 4- to 6-fold
decrease in the qEresponse at light-saturated LEF (Fig. 1A). In
this regard, it was striking that the pgr5 mutant exhibited lowered
LEF without a corresponding increase in qE sensitivity, in
contrast to what was observed in the Wt upon lowering CO2
Fig. 3 shows that gH?decreased in the Wt upon lowering CO2
but remained similar or was substantially increased in pgr5, as is
especially evident at the higher light intensities. Within the noise
level, plots of qEagainst pmfLEFfor Wt under the two CO2levels
and pgr5 overlapped (Fig. 4), indicating that, as was reported in
refs. 17 and 18, changes in gH?could predominantly account for
the differences in the qEresponse. We thus conclude that in pgr5
more facile proton efflux from the lumen through the ATP
synthase, accompanied by decreases in LEF and probably CEF1,
In principle, gH?could be modulated by changing the
specific activity of ATP synthase or its content in the thyla-
koids. Hence, a ?2-fold increase in the size of the ATP
synthase pool could give rise to the observed ?2-fold increase
(i.e., at higher light intensities) in gH?in pgr5 (Fig. 3).
However, ATP synthase content in Wt and pgr5 was estimated
by Western analyses and found to be essentially identical (Fig.
4 Inset). In addition, low light-induced activation of the ATP
synthase by thioredoxin and leakage of the thylakoid mem-
brane to protons were indistinguishable between Wt and pgr5
(data not shown), essentially as seen for other C3plants (38).
These data, taken together with the observed similarities in
gH?at low light, lead us to conclude that the differences in gH?
between Wt and pgr5 were caused by alterations in steady-state
substrate or affecter concentrations (17).
The decrease in maximal LEF in pgr5 is probably due to loss
(31, 32). A similar decrease in LEF was seen when CO2 was
lowered, but in contrast to the enhanced gH?that occurred in the
absence of PGR5, such a decrease in LEF was accompanied by
substantial decreases in gH?(Fig. 3), resulting in a net increase
in both pmf and qE. These results demonstrate an important role
that regulates light capture (39). Excessive turnover rates (i.e.,
large gH?values) will result in facile proton efflux, preventing
buildup of pmf and diminishing the qEresponse. On the other
hand, inappropriate decreases in ATP synthase turnover rates
can result in excessive buildup of pmf, over-acidifying the lumen
and causing subsequent pH-induced degradation of the photo-
synthetic apparatus (4, 40).
From the above, we conclude that changes in CEF1 upon loss
of PGR5 constitute a flux of protons approximately ?13% of
that from LEF, resulting in a commensurate decrease in ATP
output. Because consumption of ATP and NADPH by the
Calvin–Benson cycle is coupled, even a small ATP?NADPH
imbalance could conceivably give rise to not only a buildup of
ADP and [Pi], but also a substantial reduction of NADP?,
restricting the availability of PSI electron acceptors and, thereby,
lowering LEF, as was observed in pgr5 both here and in ref. 31.
Possible Causal Relation Between Pgr5?and gH?. We proposed in
ref. 17 that lowering CO2will lead to the buildup of phosphor-
ylated metabolites in the stroma, depleting stromal [Pi] below its
KM(?1 mM) at the ATP synthase. This reaction will result in
lowering of the effective gH?and subsequent increases in
steady-state pmf and qE. A small ATP?NADPH imbalance is
expected to result from the absence of the PGR5-mediated
CEF1. The deficit is obviously satisfied but only by substantially
slower processes, e.g., alternative cyclic electron transfer pro-
cesses or export of NADPH (12, 16). We thus expect in pgr5 a
buildup of stromal [Pi] above its KM at the ATP synthase,
maintaining high gH?even when LEF is restricted. Thus, in this
model, the loss of CEF1 in pgr5 indirectly attenuates both
steady-state pmf and qE.
These results support a ‘‘division of labor’’ model for pmf
synthase (gH?). Estimates of gH?in Wt and pgr5 from 26 to 216 ?mol photons
during a 300-ms dark perturbation of steady-state conditions. Conditions and
symbols are as in Fig. 1. Error bars represent SE for n ? 3–6.
The light intensity dependence of the proton conductivity of the ATP
the pmf generated solely by LEF. Estimates of energy-dependent quenching
1 and 2, respectively. ATP synthase content in Wt (A) and pgr5 (B) was
1. Error bars represent SE for n ? 3–6.
The relationship between energy-dependent exciton quenching and
www.pnas.org?cgi?doi?10.1073?pnas.0503952102 Avenson et al.
modulation, whereby Type I mechanisms act mainly to adjust
ATP?NADPH output, whereas Type II mechanisms alter the
sensitivity of antenna regulatory pathways while maintaining
pmf in an optimal range for energy transduction. Finally, it is
clear from these results that a further understanding of the
interaction of the photosynthetic apparatus within the plant will
require an integrated, yet quantitative, ‘‘systems’’ approach on
the intact plant under true steady-state conditions. Spectro-
scopic tools, such as we have applied here, will be essential for
This work was supported by U.S. Department of Energy Grant DE-
FG03-98ER20299 and U.S. National Science Foundation Grant IBN-
1. Ort, D. R. & Yocum, C. F. (1996) in Oxygenic Photosynthesis: The Light
Reactions, ed. Yocum, C. F. (Kluwe, Dordrecht, The Netherlands), pp. 1–9.
2. Kramer, D. M., Cruz, J. A. & Kanazawa, A. (2003) Trends Plant Sci. 8, 27–32.
3. Cruz, J. A., Sacksteder, C. A., Kanazawa, A. & Kramer, D. M. (2001)
Biochemistry 40, 1226–1237.
4. Kramer, D., Sacksteder, C. & Cruz, J. (1999) Photosynth. Res. 60, 151–163.
5. Capaldi, R. A. & Aggeler, R. (2002) Trends in Biochem. Sci. 27, 154–160.
6. Fischer, S. & Gra ¨ber, P. (1999) FEBS Lett. 457, 327–332.
7. Asada, K. (2000) Philos. Trans. R. Soc. London B 355, 1419–1431.
8. Muller, P., Li, X. & Niyogi, K. K. (2001) Plant Physiol. 125, 1558–1566.
9. Niyogi, K. K. (1999) Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 333–359.
10. Niyogi, K. K. (2000) Curr. Opin. Plant Biol. 3, 455–460.
11. Li, X., Bjorkman, O., Shih, C., Grossman, A. R., Rosenquist, M., Jansson, S.
& Niyogi, K. K. (2000) Nature 403, 391–395.
12. Kramer, D. M., Avenson, T. J. & Edwards, G. E. (2004) Trends Plant Sci. 9,
13. Allen, J. F. (2002) Cell 110, 273–276.
14. Allen, J. F. (2003) Trends Plant Sci. 8, 15–19.
15. Nixon, P. J. & Mullineaux, C. W. (2001) in Advances in Photosynthesis and
Respiration: Regulation of Photosynthesis, eds. Aro, E. & Anderson, B. (Kluwer,
Dordrecht, The Netherlands), Vol. 11.
16. Cruz, J. A., Avenson, T. J., Kanazawa, A., Takizawa, K., Edwards, G. E. &
Kramer, D. M. (2004) J. Exp. Bot. 56, 395–406.
17. Kanazawa, A. & Kramer, D. M. (2002) Proc. Natl. Acad. Sci. USA 99,
18. Avenson, T. J., Cruz, J. A. & Kramer, D. M. (2004) Proc. Natl. Acad. Sci. USA
19. Heber, U. & Walker, D. (1992) Plant Physiol. 100, 1621–1626.
20. Demmig-Adams, B. & Adams, W. W. I. (1992) Annu. Rev. Plant Physiol. Plant
Mol. Biol. 43, 599–626.
21. Joe ¨t, T., Genty, B., Josse, E. M., Kuntz, M., Cournac, L. & Peltier, G. (2002)
J. Biol. Chem. 277, 31623–31630.
22. Aluru, M. R. & Rodermel, S. R. (2004) Physiol. Plant. 120, 4–11.
23. Johnson, G. N. (2004) Trends Plant Sci. 9, 570–571.
24. Kramer, D. M., Avenson, T. J. & Edwards, G. E. (2004) Trends Plant Sci. 9,
Acad. Sci. USA 97, 14283–14288.
26. Sacksteder, C. & Kramer, D. M. (2000) Photosynth. Res. 66, 145–158.
27. Kramer, D. & Sacksteder, C. A. (1998) Photosynth. Res. 56, 103–112.
28. Avenson, T. J., Kanazawa, A., Cruz, J. A., Takizawa, K., Ettinger, W. E. &
Kramer, D. M. (2005) Plant Cell Environ. 28, 97–109.
29. Witt, H. T. (1979) Biochim. Biophys. Acta 505, 355–427.
30. Genty, B., Briantais, J.-M. & Baker, N. R. (1989) Biochim. Biophys. Acta 990,
31. Munekage, Y., Hojo, M., Meurer, J., Endo, T., Tasaka, M. & Shikanai, T.
(2002) Cell 110, 361–371.
32. Munekage, Y., Hashimoto, M., Miyake, C., Tomizawa, K., Endo, T., Tasaka,
M. & Shikanai, T. (2004) Nature 429, 579–582.
33. Golding, A. J. & Johnson, G. N. (2003) Planta 218, 107–114.
34. Sacksteder, C. A., Jacoby, M. E. & Kramer, D. M. (2001) Photosynth. Res. 70,
35. Donahue, R. A., Poulson, M. E. & Edwards, G. E. (1997) Photosynth. Res. 52,
36. Jauh, G. Y., Phillips, T. E. & Rogers, J. C. (1999) Plant Cell 11, 1867–
37. Kramer, D. M. & Crofts, A. R. (1996) in Photosynthesis and the Environment.
Advances in Photosynthesis, ed. Baker, N. (Kluwer, Dordrecht, The Nether-
lands), pp. 25–66.
38. Kramer, D. & Crofts, A. (1989) Biochim. Biophys. Acta 976, 28–41.
39. Herbert, S. K. (2002) Proc. Natl. Acad. Sci. USA 99, 12518–12519.
40. Majeran, W., Olive, J., Drapier, D., Vallon, O. & Wollman, F. A. (2001) Plant
Physiol. 126, 421–433.
Avenson et al.
July 5, 2005 ?
vol. 102 ?
no. 27 ?