Plant Physiol. (1998) 118: 103-113
pH-Dependent Photosystem II Fluorescence Quenching Induced by
Saturating, Multiturnover Pulses in Red Algae1
Estelle Delphin,
Jean-Claude Duval,
Anne-Lise Etienne, and
Diana Kirilovsky*
Photorégulation et Dynamique des Membranes
Végétales, Unité de Recherche Associée 1810, Centre National de la Recherche Scientifique, École Normale
Supérieure, 46 rue d'Ulm, 75230 Paris cedex 05, France
 |
ABSTRACT |
We have previously shown that in the
red alga Rhodella violacea, exposure to continuous low
intensities of light 2 (green light) or near-saturating intensities of
white light induces a 
pH-dependent PSII fluorescence quenching. In
this article we further characterize this fluorescence quenching by
using white, saturating, multiturnover pulses. Even though the pulses
are necessary to induce the pH and the quenching, the development of
the latter occurred in darkness and required several tens of seconds.
In darkness or in the light in the presence of
2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone, the
dissipation of the quenching was very slow (more than 15 min) due to a
low consumption of the pH, which corresponds to an inactive ATP
synthase. In contrast, under far-red illumination or in the presence of
3-(3,4-dichlorophenyl)-1,1
-dimethylurea (only in light), the
fluorescence quenching relaxed in a few seconds. The presence of
N,N-dicyclohexyl carbodiimide hindered this relaxation. We propose
that the quenching relaxation is related to the consumption of pH by
ATP synthase, which remains active under conditions favoring
pseudolinear and cyclic electron transfer.
| |
INTRODUCTION |
Photosynthetic organisms convert light energy into chemical
energy. The energy absorption occurs at the level of the antenna of two
membrane-pigment complexes, PSI and PSII. The absorbed energy is
transferred to the reaction centers, which operate in series. The
electron transport from water to NADP+ is coupled
to a proton translocation across the membrane, generating a proton
motive force for the synthesis of ATP.
Trapping by the photochemical centers is the main pathway of
deactivation of the excitons formed by the absorption of photons. Fluorescence and thermal dissipation are the other ways to deactivate these excitons. Photochemistry and qP are maximal
when all PSII centers are open and the 
o is
low. When all of the centers are closed, qP is
suppressed and the o is concomitantly
increased to its maximal value (m).
Fluorescence is modulated by the oxidoreduction state of the primary
acceptor QA (Duysens and Sweers, 1963
; van Gorkom, 1974). Nonphotochemical processes can also reduce the o. Several mechanisms may be involved at the
level of the antenna system and the PSII reaction centers (Krause and
Weis, 1991; Horton and Ruban, 1992). NPQ of fluorescence is ascribed to
three major processes: state transitions, qE,
and photoinhibition. These different mechanisms have been widely
studied either in vitro, in thylakoid and chloroplast, or in vivo, in
leaves and green algae cells. The different types of NPQ can be
recognized because they are developed at different light intensities,
inhibited by specific chemical treatments, and relaxed in the dark with
characteristic kinetics (Demmig and Winter, 1988; Horton and Hague,
1988; Quick and Stitt, 1989; Lee et al., 1990; Walters and Horton,
1991, 1993).
qE is related to a transthylakoid proton gradient
formed during electron transport (Krause, 1973; Briantais et al., 1980; Krause et al., 1982). It was shown that in higher plants and in green
and brown algae, part of this quenching is correlated to the
dissipation of an excess of excitation energy in the PSII antenna
complex. The formation of this quenching is accompanied by the
accumulation of de-epoxidated xanthophyll (Demmig et al., 1987; Bilger
et al., 1989; Demmig-Adams, 1990; Gilmore and Yamamoto, 1991) and
conformational changes (most probably, aggregation) of LHCII (Horton et
al., 1991, 1996; Noctor et al., 1991; Ruban et al.,
1992a, 1993). Part of the quenching was also attributed to
energy dissipation in the reaction center by radiationless decay (Weis
and Berry, 1987) or rapid charge recombination (Krieger et al., 1992;
Krieger and Weis, 1993). The mechanism of
qE concerns PSII, but not PSI.
Different incident wavelengths can selectively excite one of the two
photosystems. Higher plants, green algae, and cyanobacteria have
developed a mechanism called state transitions to optimize the
utilization of the captured energy and to regulate the production of
ATP and NADPH. In higher plants and green algae, the redistribution of
absorbed excitation energy between PSI and PSII involves
redox-dependent phosphorylation/dephosphorylation reactions of LHCII,
leading to opposite changes of the cross-section of the PSII and PSI
antenna complexes (for review, see Haworth et al., 1982; Williams and Allen, 1987; Allen, 1992, 1995). Both photosystems are involved in this
mechanism.
In red algae and cyanobacteria, the energy collection of PSII is mainly
achieved by the phycobilisomes, large extramembrane complexes formed by
phycobiliproteins, whereas Chl a is the major energy-collector pigment in PSI. The mechanisms involved in PSII fluorescence quenching in phycobilisome-containing organisms are not as
well characterized as in higher plants. It was proposed that state
transitions were related to redistribution of the absorbed energy by
changes in the spillover between PSII and PSI (Ley and Butler, 1980;
Biggins et al., 1984; Bruce et al., 1985; Olive et al., 1986; Vernotte
et al., 1990) or to changes in the cross-section of the antenna of both
PSI and PSII (Allen et al., 1985; Sanders and Allen, 1987, 1988). It
was also suggested that only PSII is involved in the mechanism of
fluorescence quenching (Satoh and Fork, 1983). However, few studies
analyzed the possibility of the existence of a qE
in red algae or cyanobacteria.
Recently, we have studied the adaptation to changes in light intensity
and quality in the red alga Rhodella violacea (Delphin et
al., 1995, 1996). We demonstrated in vivo that the fluorescence changes
produced by illumination at different wavelengths were independent of
protein phosphorylation (Delphin et al., 1995). We have also shown that
illumination of R. violacea cells with light 2 (light
preferentially absorbed by the phycobiliproteins) induced a large
quenching of PSII fluorescence. This quenching was suppressed by
addition of the uncoupler NH4Cl, whereas it was
maintained when the cells were exposed to light 1 (light preferentially absorbed by the PSI) illumination in the presence of DCCD, an inhibitor
of ATP synthase. The level of the PSI-related fluorescence (detected at
low temperature) did not change under these different conditions. We
concluded that the fluorescence changes commonly associated with state
2 transition are in fact due to a pH-dependent quenching (Delphin et
al., 1996).
It has been generally assumed that the fluorescence changes induced by
different light conditions in red algae and cyanobacteria are related
to the same mechanism. However, we have demonstrated that such changes
are associated with a transmembrane pH in red algae (Delphin et al.,
1996), and Campbell and Öquist (1996) proposed that in
cyanobacteria, the changes in o are only
related to state transitions. In our previous studies on fluorescence quenching mechanisms in red algae, we used the light-saturation-pulse method, in which brief pulses of intense light remove all of the qP and reveal the remaining quenching, which is
nonphotochemical in nature, as shown by the level of the
Fm assessed during the pulse
(Bradbury and Baker, 1984; Renger and Schreiber, 1986; Schreiber et
al., 1986, 1995; Quick and Stitt, 1989; Krause and Weis, 1991; Walters
and Horton, 1991; Lokstein et al., 1993). We observed that the
saturating, multiturnover, white pulses induced a large quenching of
Fm. In this article we further
characterize the PSII fluorescence quenching mechanisms in red algae by
using saturating, multiturnover pulses at different intensities,
lengths, and frequencies.
| |
MATERIALS AND METHODS |
Culture of Algae
Red algae (Rhodella violacea) cells were grown
autotrophically in Erdschreiber medium (modified seawater) enriched
with FeCl3·6H2O (0.6 mg
L
1) at 20°C. The cultures were continuously
flushed with sterile air. Light was provided by cool-white fluorescent
tubes (Philips, Eindhoven, The Netherlands) at an intensity of 60 µmol photons m2 s1 in
a 16-h light/8-h dark cycle. Cells were collected at a concentration of
about 8.0 × 105 cells
mL1, corresponding to a Chl concentration of
5.5 µg Chl mL1. Porphyridium
cruentum cells were grown in the same conditions as R. violacea cells, except that the culture medium was artificial seawater prepared according to the method of Jones et al. (1963) and the light intensity was 30 µmol photons
m2 s1. Cells were
collected at a concentration of about 4.5 × 106 cells mL1,
corresponding to a Chl concentration of 4.5 µg
mL1. The cells were maintained under white,
low-light conditions. Each experiment began with 5 min of dark
adaptation followed by 4 min of cell incubation in far-red light.
Fluorescence Measurements
Chl o was measured at the same
temperature as that of the growth cultures with a pulse-amplitude
modulated fluorometer (model 101, Heinz-Walz, Effelrich, Germany)
adapted to an oxygen electrode (model DW1, Hansatech, King's Lynn,
UK), as previously described by Arsalane et al. (1994). In the absence
of any actinic light, the minimal o
(Fo or Fo) was
measured by the nonactinic-modulated beam. Saturating, multiturnover,
white-light pulses (3200 µmol photons m2
s1) were applied to assess the maximal
o (Fm or
Fm). In this paper the maximal
o obtained by a train of saturating white
light pulses under nonactinic-modulated light is also called
Fm. The pulses were generated by an
electronic shutter (Uniblitz, Vincent, Rochester, NY) in front
of a quartz-iodine lamp (model KL-1500, Schott, Mainz, Germany) that
was continuously on. The shutter was controlled by the accessory module
pulse-amplitude modulated fluorometer (model 103, Heinz-Walz).
Two types of recordings were carried out in these experiments, those
measured in milliseconds and those measured in seconds. For the slow
fluorescence changes, the sampling was done every 10 ms and averaged
every 3 s. For a correct evaluation of
Fm (or Fm)
during a light pulse of 800 ms, the signal was averaged over the second
half of the light pulse (from 400 to 800 ms). During inductions in the
millisecond range, the fluorescence signal was sampled every
millisecond. The Fo level was assessed with
the modulated detector beam before the shutter opened. There was a 3-ms
delay between the trigger of the signal recording and the shutter
opening. The shutter took 3 ms to be fully open.
The different background continuous illuminations were obtained from
another KL-1500 lamp, which was either used as white light or filtered
at 540 nm for green light (filter 70617 AM-4064, Oriel, Stratford, CT),
at 601 nm for orange light (filter: 40 599 10, Balzers Pseiffer, North
America, Hudson, NH), or at 435 nm for blue light (filter 4-35,
Corning Instruments, Corning, NY). Far-red light (735 ± 10 nm)
was obtained from a light-emitting diode (102-FR, Heinz-Walz). Data
acquisition, shutter control, and pulse averaging were driven by
homemade software through a 12-bit analogic digital converter, as
previously described (Arsalane et al., 1994). When indicated, 77 K
fluorescence emission spectra were measured as described in Delphin et
al. (1996).
| |
RESULTS |
Effect of Saturating, Multiturnover Pulses on Maximal Fluorescence
This study was performed on two strains of red algae,
R. violacea and P. cruentum. We have
already shown that in R. violacea, a large pH-dependent
quenching of PSII fluorescence is generated by continuous green light
(absorbed mainly by phycobilisomes) even at low light intensities
(Delphin et al., 1996). Similar results have been obtained in P. cruentum cells. In this alga, as in R. violacea, the
Fm level was maximal in far-red-adapted cells. Under green-light illumination, a large
Fm quenching was developed. The main part
of the fluorescence quenching was suppressed when the pH across the
membrane was suppressed by addition of NH4Cl or
nigericin, even under green-light illumination (data not shown).
Near-saturating and saturating intensities of continuous white light
also induce a large pH-dependent fluorescence quenching in red alga
cells (Delphin et al., 1996). Moreover, in our previous studies, we
have observed that the repetitive application of saturating pulses of
white light induces a large decrease of Fm
in red alga cells. R. violacea and P. cruentum
dark-adapted cells were preincubated in far-red light (75 µmol
photons m2 s1) for 4 min to ensure a maximal level of Fm
(Delphin et al., 1996). The far-red light was turned off before a train
of saturating pulses (800 ms, 3200 µmol photons
m2 s1) was applied. The
interpulse time was 60 s. Figure 1
shows that Fv
(Fm 
Fo)
decreased during a series of pulses to reach a level of 20% to
30% of the maximal Fv in R. violacea cells. No variation of the
Fo level was detected. This
Fv quenching was also observed when the
train of saturating pulses was directly applied to dark-adapted cells.
Similar results were obtained with P. cruentum cells (data
not shown).
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| Figure 1.
Effect of a train of saturating pulses (800 ms,
3200 µmol photons m2 s1) on
Fm and Fo
levels in R. violacea cells. Dark-adapted (for 5 min)
cells were preincubated for 4 min in far-red light (75 µmol photons
m2 s1). Far-red light was turned off before
the pulses were applied; pulses were spaced by 60 s. rel.,
Relative units.
|
|
Low-temperature fluorescence spectra of cells preadapted to far-red
light and of those exposed to a sequence of saturating pulses were
performed in the presence of an external fluorescence probe
(phycocyanin from Spirulina maxima). Low-temperature
emission spectra were deconvoluted into Gaussian components, and the
areas corresponding to F695 (PSII
fluorescence), F718, (PSI fluorescence), and F650 (probe fluorescence) were
calculated. The F695 to
F718 ratio decreased from 1.4 to 0.7 after
the sequence of saturating pulses. The calculated ratio of
F718 to F650
showed that PSI fluorescence was similar before and after the sequence
of pulses (F718 to
F650= 1 ± 0.03). In contrast, the
F695 to F650
ratio varied from 1.4 ± 0.04 to 0.8 ± 0.03, indicating a
decrease in PSII fluorescence. These results suggest that a specific
quenching of F695 was responsible for the
decrease of the F695 to
F718 ratio.
Effect of the Intensity and Length of Light Pulses
To further characterize the effect of individual pulses on the
development of fluorescence quenching, the length and intensity of the
pulses were systematically varied. To monitor the fluorescence changes
occurring during the pulse, induction curves were recorded during the
pulse at different intensities: 15, 50, 300, and 3200 µmol photons
m2 s1 (Fig.
2A). The pulse was applied to cells
preincubated for 4 min in far-red light. The induction curves were
recorded 30 s after the far-red light was turned off. Under light
intensities higher than 300 µmol photons m2
s1 during the pulse, the P level was the
maximum o attained during the transition.
Under 3200 µmol photons m2
s1 light intensity, the P level was attained
after 70 ms (trace 4), whereas under 300 µmol photons
m2 s1, the P level was
reached in 300 ms (trace 3). When the light intensity was lower than
300 µmol photons m2
s1, the P level was not reached (traces 2 and
1, Fig. 2A). Fluorescence quenching was not developed during the light
pulse. Figure 2B shows that the P level was slightly higher than the
Fm measured in the presence of
105 M DCMU. In the presence of DCMU, the
oxidized PQ pool is a quencher of fluorescence (Vernotte et al.,
1979).
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| Figure 2.
Fluorescence inductions of R. violacea cells. A, 800-ms duration pulses of varying
white-light intensities: 15 (trace 1), 50 (trace 2), 300 (trace 3), and
3200 (trace 4) µmol photons m2 s1; B, a
saturating pulse (800 ms, 3200 µmol photons m2
s1) in the absence or in the presence of DCMU (10 µM); C, a train of saturating pulses (800 ms/300 µmol
photons m2 s1) spaced by 60 s.
Fluorescence inductions during the 1st, 2nd, 3rd, 4th, and 8th pulse
are presented. Dark-adapted cells were preilluminated with far-red
light (75 µmol photons m2 s1) for 4 min.
Induction curves were recorded 30 s after far-red light was
switched off.
|
|
Figure 2C shows the fluorescence induction generated during the 1st,
2nd, 3rd, 4th, and 8th pulses of 300 µmol photons
m2 s1 and 800 ms in
duration. Each additional pulse produced an additional decrease of the
variable fluorescence visible during the subsequent pulse. We observed
that almost all of the levels of fluorescence were affected by
quenching. However, the P level was the most affected: at the end of a
series of pulses the fast initial phase was diminished by one-half,
whereas the slower phase (rise to P level) almost disappeared. The
Fo remained constant.
The quenching of the variable fluorescence induced by the first light
pulse 800 ms in duration and detected by the second pulse was
proportional to the light intensity (Fig.
3A). If the length of pulses was varied
while fixing the intensity to 300 µmol photons
m2 s1, we observed that
the quenching increased with the length of the pulse up to 400 ms (Fig.
3B). As shown in Figure 3C, 300 ms was the time required to reach the P
level. No additional quenching was induced if pulses were of a longer
duration (up to 800 ms).
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| Figure 3.
Effect of the intensity and the length of the
pulse on the extent of the fluorescence quenching in R. violacea cells. A, Pulses of 800 ms at different light
intensities. B, Pulses of different lengths at 300 µmol photons
m2 s1. Calculations of the percent of
quenching produced by a first pulse were done on the variable
fluorescence (Fv = Fm Fo) of a
second pulse (800 ms, 3200 µmol photons m2
s1) applied 60 s after the first pulse. A pulse
given after 2 min of far-red illumination was set as 100% of
Fv.
|
|
Fv Quenching and the Dark Period
between the Pulses
To determine the kinetics of formation and relaxation of
Fv quenching, the duration of dark periods
between pulses was varied. A train of saturating pulses (800 ms, 3200 µmol photons m2 s1)
spaced by 10, 30, 60, 200, 300, or 600 s was applied to far-red light-adapted cells after far-red light was turned off. In both strains
and for all the frequencies of pulses, the level of
Fv decreased (Fig.
4, A and B).
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| Figure 4.
Effect of the pulse frequency on the
Fv level in far-red-light-adapted R. violacea cells (A) and P. cruentum cells (B) as
a function of the number of pulses. Pulses of 800 ms, 3200 µmol
photons m2 s1, were spaced by 10, 30, 60, 200, 300, or 600 s. A pulse given after 2 min of far-red
illumination was set as 100% of Fv. C,
Kinetics of fluorescence quenching during dark incubation after a light
pulse (800 ms, 3200 µmol photons m2 s1)
in R. violacea ( ) and P. cruentum
cells ( ). The detection and calculation of the percent of quenching
was done as in Figure 3 except that the detecting pulse was applied
after different times.
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|
Figure 4C shows the quenching of Fv
induced by one pulse and detected by a second pulse after different
dark periods. A dark period between 100 and 200 s was optimal to
reach the quenched state in R. violacea. When the dark
period was longer than 200 s, the effect of the pulse began to be
dissipated between the pulses (Fig. 4, A and C). The effect of each
pulse seemed to dissipate faster in P. cruentum than in
R. violacea cells. In P. cruentum cells a dark
period of 30 s was optimal to reach a maximal
Fm quenching (Fig. 4, B and C). When the
spacing between the pulses was 60 s or more, a reverse of the
quenching became evident.
Origin of the Fm Quenching Induced by
Saturating White Pulses
NH4Cl Effect
As mentioned above, the formation of a pH across the thylakoid
membrane is responsible for the fluorescence quenching observed under
continuous, near-saturating white light (Delphin et al., 1996).
Therefore, we assumed that the quenching induced by saturating white
pulses would be pH dependent. To confirm this hypothesis, chemical
and light treatments that can alter the extent of the pH were
applied before or during the pulse sequences. In the first experiment,
the pH was not allowed to accumulate during the series of saturating
pulses: the uncoupler NH4Cl (2 mM)
was added 1 min before the first pulse. Figure
5 shows that in the presence of the
uncoupler, repetitive pulses did not affect (P. cruentum
cells) or slightly affected (R. violacea cells) the level of
Fm (or Fv).
In the second experiment, NH4Cl was added after 12 pulses, after the fluorescence quenching was formed. Addition of the
uncoupler suppressed most of the quenching (Fig. 5). The same result
was obtained when the uncoupler nigericin was used instead of
NH4Cl (data not shown).
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| Figure 5.
Effect of NH4Cl on the
Fv level in R. violacea
(circles) and P. cruentum (triangles) cells during a
train of saturating pulses (800 ms, 3200 µmol photons
m2 s1) spaced by 60 s.
NH4Cl (2 mM) was added 1 min before the onset
of the train of pulses (open symbols), or after 14 pulses (closed
symbols). Calculations of the percentage of quenching were done on
the variable fluorescence recorded during pulses,
Fv = Fm Fo. The first pulse of the train was set as
100% of Fv.
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|
Far-Red Light Effect
Because the effect of the pulses was accumulative, we assumed that
the pH was not (or not sufficiently) dissipated in darkness between
the pulses. In contrast, when the pulses were separated by 60 s of
far-red illumination, only a slight decrease of
Fm was observed (Fig.
6B), indicating that illumination by
far-red light favored the consumption of pH between the pulses. When DCCD (40 µM), an inhibitor of ATP synthase, was present
while the background of far-red illumination was maintained, a large quenching of Fm appeared again (Fig. 6D),
indicating the absence of pH consumption. Under modulated
illumination the decrease of Fv was
slightly faster in the presence of DCCD (Fig. 6C) than in its absence
(Fig. 6A). Under far-red illumination and in the presence of DCCD, the
decrease of Fv was even faster (Fig. 6D). These results suggest that ATP synthase was active during part of the
pulse and during far-red illumination.
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| Figure 6.
Effect of far-red light in the dissipation of
fluorescence quenching in R. violacea cells and the
influence of DCCD and DBMIB. The far-red preincubation light was
maintained during the trains of pulses in B, D, E, and F. The trains of
pulses (800 ms, 3200 µmol photons m2 s1)
were applied in the absence of any chemical (A and B) or in the
presence of DCCD (40 µM) (C and D), DBMIB (10 µM) (E), or DBMIB (10 µM) plus nigericin
(100 µM) (F). DCCD was added in the dark and incubated
for 10 min before the far-red light was turned on. DBMIB and
nigericin were added just before turning on the far-red light.
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Figure 6B shows that illumination by far-red light eliminated the
fluorescence quenching induced by the white, saturating pulses.
However, cyclic electron transport functioning under far-red illumination should induce a proton translocation across the membrane, increasing the pH. The existence of a pH coupled to cyclic
electron transport around the PSI induced by far-red illumination was
demonstrated by incubating the cells in the presence of DCCD (40 µM) under far-red illumination in the absence of light
pulses. Under these conditions, Fv
decreased 40% after 10 min and 70% after 20 min of far-red
incubation. In the absence of DCCD, no fluorescence quenching was
detected, suggesting that the pH formed was small and/or that it was
rapidly consumed.
The results presented in this section confirm that the persistence of
the fluorescence quenching in darkness was due to the inactivation of
ATP synthase in the dark, leading to a very slow consumption of the
pH. We propose that far-red light inhibited the development of
fluorescence quenching by maintaining the ATP synthase active between
the pulses. Under far-red illumination, PSI is active and it can be
assumed that reduction of PQ molecules via a NAD(P)H-PQ oxydoreductase
would provide some electrons to maintain pseudolinear electron transfer
and thioredoxin reduction, favoring ATP synthase activity.
DBMIB Effect
We assumed that DBMIB, which inhibits linear and cyclic electron
transport by interaction with the Qo site of the
cyt b6/f complex, may
hinder the ATPase activity under far-red illumination. The presence of
DBMIB (10 µM) when the train of pulses was
superimposed to far-red illumination also induced the
Fm quenching (Fig. 6D). The fact that the
Fo level between the pulses remained low
and constant indicated that the plastoquinol molecules were reoxidized during the 60-s far-red illumination between the light pulses. This
reoxidation had to occur via a pathway that did not involve the cyt
b6/f, probably via
equilibrium with molecular oxygen. Our results also indicated that the
reoxidation of the PQ molecules allowed PSII turnover during each
successive pulse. During each light pulse, even in the presence of
DBMIB, there was enough proton pumping associated with PQ reduction to
induce fluorescence quenching that was not dissipated between the
pulses. The DBMIB effect was suppressed by addition of uncouplers,
confirming that DBMIB hindered the consumption of the pH under
far-red light (Fig. 6E).
DCMU Effect
We have already reported that addition of DCMU suppressed the
fluorescence quenching induced by continuous green light (Delphin et
al., 1996). In certain conditions, DCMU is also able to eliminate the
fluorescence quenching induced by the train of pulses. Figure 7, A through C, shows the effect of DCMU
(10 µM) under different light conditions. DCMU was added
to the "quenched" sample and then the cells were illuminated for 4 min by "nonactinic" modulated light (Fig. 7A), 50 µmol photons
m2 s1 white light (Fig.
7B), and 200 µmol photons m2
s1 white light (Fig. 7C) before a train of
saturating pulses was applied. The rate of quenching relaxation
depended on the light intensity: 8 min under the modulated light (Fig.
7A), 4 min at 50 µmol photons m2
s1 of white light (Fig. 7B), and 1 min at 200 µmol photons m2 s1
(Fig. 7C). Illumination was necessary for the DCMU effect (Fig. 7D).
DCMU added in the dark, after the formation of
Fm quenching, did not suppress the
quenching even if the dark period was for 4 min (Fig. 7D). Only after
15 min of dark incubation in the presence of DCMU did the quenching
begin to relax (data not shown). This clearly rules out the possibility
of DCMU acting as an uncoupler.
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| Figure 7.
Effect of DCMU on Fm
quenching in R. violacea cells. Far-red-adapted cells
were exposed to a train of pulses after turning off the far-red light.
DCMU (10 µM) was added after six light pulses (800 ms,
3200 µmol photons m2 s1) spaced by
60 s, under the nonactinic-modulated light (0.046 µmol photons
m2 s1) (A-C) or in the dark (D). After
DCMU addition, the cells were incubated for 4 min in modulated light
(A); 50 µmol photons m2 s1 white light
(B); 200 µmol photons m2 s1 white light
(C); or darkness (D). Then, the cells in the dark were shifted to
modulated light. The other samples were maintained under the same light
conditions and a train of saturating pulses was applied.
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|
Fluorescence quenching was suppressed by DCMU in the absence but not in
the presence of DCCD (40 µM) (Fig.
8). Moreover, the presence of DCCD
largely decreased the actinic effect of the modulated light observed in
the presence of DCMU. Since DCCD is an inhibitor of ATP synthase, one
can conclude that under illumination, the presence of DCMU eliminated
the quenching by preventing the formation of the pH coupled to PSII
activity without inhibiting the consumption of the pH formed by the
illumination period preceding DCMU addition. Under illumination in the
presence of DCMU, PSI is active and pseudolinear, and cyclic electron
transport would occur. Under these conditions the pH consumption
seems to be faster than its buildup induced by cyclic electron
transport or by far-red illumination.
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| Figure 8.
Effect of DCMU in the absence or presence of DCCD
in R. violacea cells. Far-red-adapted cells were exposed
to a train of pulses in the absence (A) or in the presence (B) of DCCD
(40 µM). DCCD was added in the dark and incubated for 10 min before cells were exposed to far-red light (2 min). Far-red light
was turned off before the train of pulses was applied. DCMU (10 µM) was added in the dark and after six light pulses
spaced by 60 s. After 4 min of dark incubation the modulated light
was turned on. One pulse was immediately applied and a second one was
applied after 5 min.
|
|
| |
DISCUSSION |
We have previously shown that in red algae exposed to continuous
illumination, low and high intensities of light 2 (green light) and
near-saturating and saturating intensities of white light induced a
PSII fluorescence quenching triggered by the formation of a
transthylakoid pH and not by the reduction of the PQ. Quenching was
relaxed under conditions that suppressed the pH across the membrane,
even when the PQ remained reduced (Delphin et al., 1996). In the
present paper we demonstrated that in red algae (R. violacea and P. cruentum), white-light pulses of a wide range of
intensities and lengths were able to induce a pH-dependent
Fm quenching with no change in
Fo. When the formation of pH was
hindered by the presence of NH4Cl or nigericin,
no fluorescence quenching occurred. The quenching induced by a train of
pulses rapidly disappeared after addition of uncouplers.
Even though fluorescence quenching was induced by light, it did not
develop during a pulse with a duration less than or equal to 800 ms.
Fluorescence quenching developed during the dark period following the
pulse, indicating that, once induced, light was not necessary for its
development. The quenching was revealed by the following light pulse.
Several tens of seconds were required to develop the maximal
quenching induced by each pulse (30 s in P. cruentum and
100 s in R. violacea). For this reason, in R. violacea, pulses spaced by 100 s of darkness seemed to have a greater effect than pulses spaced by 10 s of darkness. These
results also suggest that it is not the formation of the proton
gradient that limits the rate of quenching formation. A slower process after the pulse is required for the full occurrence of the quenching. In darkness the dissipation of quenching was very slow; it took up to 5 min in P. cruentum and up to 10 min in R. violacea.
As expected, under a large range of intensities and lengths,
fluorescence quenching increased with the quanta absorbed by the cells.
However, pulses of very different lengths (70 and 800 ms for the 3200 µmol photons m2 s1
pulses or 300 and 800 ms for the 300 µmol photons
m2 s1 pulses) induced a
similar Fm quenching. To explain this
result, we propose that it takes more time to activate ATP synthase
than to reduce the PQ pool. During the first period of illumination, corresponding mainly to PQ reduction, the pH is efficiently formed by PSII and little is consumed. The time needed for the full reduction of the PQ pool depends on the light intensity of the pulse (70 ms for
the 3200 µmol photons m2
s1 pulse and 300 ms for the 300 µmol photons
m2 s1 pulse, see Fig.
5A). When the PQ pool becomes mostly reduced, the PSII activity and the
proton translocation decrease, whereas the ATP synthase is activated.
The fact that the decrease of Fv was
slightly faster in the presence of DCCD than in its absence also
suggests a small ATP synthase activity during the pulse (Fig. 6).
Our results suggest that the persistence of the fluorescence quenching
in the dark corresponds to a low consumption of the pH. As soon as
the pH was nullified, the maximal fluorescence was restored. In
higher plants the activity of ATP synthase depends on the pH and on
the redox state of the thioredoxin (Vallejos et al., 1983; Junesch and
Gräber, 1984; Mills and Mitchell, 1984): The reduced thioredoxin
activates ATP synthesis by reduction of the 
-subunit of ATP synthase
(Nalin and McCarty, 1984). We propose that the same is true in red
algae. In dark-adapted cells the ATP synthase seems to be inactive;
upon illumination it is activated and the rate of pH consumption
slowly increases. In the dark it rapidly becomes inactive. The
accumulative effect of pulses can be explained by the very slow
consumption of the pH during the dark periods in red algae cells.
Cyclic electron transfer occurring under far-red light induced a pH
gradient sufficiently large to generate fluorescence quenching even in
the absence of white light pulses. However, this quenching was formed
only in the presence of DCCD, an inhibitor of ATP synthase. In
contrast, in the absence of DCCD, this quenching was not observed, suggesting that the consumption of the pH was faster than its formation. Alternatively, it can be assumed that the formation of the
NPQ is more sensitive to H+ domains localized
around PSII than to the pH across the membrane generated by cyclic
electron transfer. Our results also suggest that far-red illumination
maintained ATP synthase in an active state. Under far-red illumination,
thioredoxin could be reduced via pseudolinear electron transfer allowed
by the PQ reduction by a NAD(P)H-PQ oxidoreductase. In this case, the
thioredoxin reduction and the pH gradient generated by cyclic electron
transport could contribute to maintain ATP synthase activity. Moreover, the far-red-generated pH seemed to be sufficient to maintain the
activity of ATP synthase between the pulses. When the pulses were
applied in the presence of far-red light, almost no fluorescence quenching was observed between the light pulses, indicating that the
pH was consumed (Fig. 6). This hypothesis was supported by the fact
that in the presence of DCCD the fluorescence quenching developed.
The antagonistic effects of DCMU and DBMIB can be interpreted in terms
of their effects on pH but not in terms of their effects on the
redox state of PQ. DCMU addition relaxed fluorescence quenching under illumination, whereas it had no effect in the presence of DCCD or in darkness. In the presence of DCMU and light, as was the case
under far-red illumination, cyclic and pseudolinear electron transport
via PSI were active. We propose that these conditions maintained ATP
synthase activity, enabling a rapid pH consumption and avoiding
quenching formation or relaxing the quenching previously induced.
Addition of DBMIB, which inhibited both cyclic and pseudolinear electron transport, suppressed the effect of far-red illumination. Since the presence of uncouplers annulled the effect of DBMIB, we
conclude that this chemical indirectly decreased ATP synthase activity
by inhibiting pseudolinear and cyclic electron transport.
It has been demonstrated previously that, in higher plants, DCCD, in
addition to inhibiting the ATP synthase, inhibits the H+ release into the lumen following water
splitting, hindering pH (Jahns et al., 1988) and
qE formation (Ruban et al., 1992b). These effects of DCCD are due to modifications of amino acids on LHCII polypeptides (Jahns et al., 1988; Ruban et al., 1992b; Walters et al., 1996). In red algae DCCD's effect on qE
seems to be only related to the maintenance of a proton gradient by
inhibition of the ATP synthase. Addition of uncouplers such as
NH4Cl or nigericin annulled the effect of DCCD
(data not shown). However, our results do not allow us to clearly
distinguish between the effect on qE of a local
concentration of H+ around PSII to that of a
nonlocalized pH across the membrane. We demonstrated that conditions
favoring PSII activity (green and white light [Delphin et al., 1996;
this article]) induce a pH-dependent quenching, whereas under
conditions favoring cyclic electron transfer (far-red illumination,
presence of DCMU), no fluorescence quenching was observed in the
absence of DCCD. These results favor the hypothesis involving a
localized proton gradient around the PSII on the induction of
qE in red algae.
Contrasting hypotheses have been put forward to explain the molecular
mechanism of qE. In most studies in leaves and
chloroplasts, a decrease of Fo has been
associated with Fm quenching (Bilger and
Schreiber, 1986; Rees et al., 1990; Walters and Horton, 1991; Gilmore
and Yamamoto, 1992; Horton and Ruban, 1993). This feature reflects the
energy de-excitation in the PSII antenna, which decreases the lifetime
of excited Chls (Horton and Ruban, 1992; Walters and Horton, 1993;
Horton et al., 1996). Another hypothesis for the mechanism of
qE in higher plants is the energy dissipation in
PSII reaction centers. It was proposed that the acidification of the
thylakoid lumen provokes the release of Ca2+,
inhibiting the electron donation to the reaction center II and resulting in quenching either through rapid-charge recombination or by
the accumulation of a "quencher" such as
P680+ (Weis and Berry, 1987; Krieger et al.,
1992; Krieger and Weis, 1993). In red algae no change in the minimal
fluorescence was detected under the different light regimes used (train
of saturating pulses [this study], continuous low light 2, or
near-saturating white light illumination [Delphin et al., 1996]),
suggesting that PSII fluorescence quenching might occur in the reaction
center and not in the antenna. Saturation curves of oxygen evolution also suggested that photon flux to the centers is similar in the absence or presence of quenching (Delphin et al., 1996). However, preliminary measurements of oxygen emission per flash in a Joliot-type electrode suggest that, under saturating light intensities, oxygen yield per flash and therefore the number of active PSII centers are not
affected by fluorescence quenching. Further studies are necessary to
elucidate the molecular mechanism of fluorescence quenching in red
algae.
| |
FOOTNOTES |
1
This work was supported by a grant from the
Société de Secours des Amis des Sciences to E.D.
*
Corresponding author; e-mail kirilov{at}wotan.ens.fr; fax
33-1-44-32-39-35.
Received February 18, 1998;
accepted May 29, 1998.
| |
ABBREVIATIONS |
Abbreviations:
Chl, chlorophyll.
DBMIB, 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone.
DCCD, N,N-dicyclohexyl carbodiimide.
Fo, Fm, and Fv,
initial, maximum, and variable Chl fluorescence in dark-adapted
samples, respectively .
Fo, Fm, and Fv,
initial, maximum, and variable Chl fluorescence in illuminated samples,
respectively .
LHCII, light-harvesting Chl
a/b protein.
NPQ, nonphotochemical
quenching.
o, fluorescence yield.
PQ, plastoquinone.
qE, energy-dependent quenching.
qP, photochemical quenching.
| |
ACKNOWLEDGMENTS |
We thank G. Paresys and J.-P. Roux for their excellent computer
and electronic assistance and for writing the computer software used in
the pulsed-amplitude fluorimeter.
| |
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Abstract
Introduction
