Laboratoire Organismes Photosynthétiques et Environnement,
Unité Mixte de Recherche-Centre National de la Recherche
Scientifique 8543, Ecole Normale Supérieure, 46 rue
d'Ulm, 75230 Paris cedex 05, France (J.L., B.R., A.-L.E.); and
Department of Biophysics, Huygens Laboratory, Leiden University, P.O.
Box 9504, 2300 RA Leiden, The Netherlands (H.J.v.G.)
The pool size of the xanthophyll cycle pigment
diadinoxanthin (DD) in the diatom Phaeodactylum
tricornutum depends on illumination conditions during culture.
Intermittent light caused a doubling of the DD pool without significant
change in other pigment contents and photosynthetic parameters,
including the photosystem II (PSII) antenna size. On exposure to
high-light intensity, extensive de-epoxidation of DD to diatoxanthin
(DT) rapidly caused a very strong quenching of the maximum chlorophyll
fluorescence yield (Fm, PSII reaction centers closed), which was fully reversed in the dark. The
non-photochemical quenching of the minimum fluorescence yield
(Fo, PSII centers open) decreased the
quantum efficiency of PSII proportionally. For both
Fm and Fo, the
non-photochemical quenching expressed as
F/F' 
1 (with F' the
quenched level) was proportional to the DT concentration. However, the
quenching of Fo relative to that of
Fm was much stronger than random quenching
in a homogeneous antenna could explain, showing that the rate of
photochemical excitation trapping was limited by energy transfer to the
reaction center rather than by charge separation. The cells can
increase not only the amount of DT they can produce, but also its
efficiency in competing with the PSII reaction center for excitation.
The combined effect allowed intermittent light grown cells to
down-regulate PSII by 90% and virtually eliminated photoinhibition by
saturating light. The unusually rapid and effective photoprotection by
the xanthophyll cycle in diatoms may help to explain their dominance in
turbulent waters.
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INTRODUCTION |
Photosynthetic organisms have
developed strategies to optimize light harvesting at low intensities
while minimizing photoinhibitory damage due to excess energy at
high-light intensity. From hours onwards, they regulate the quantity
and composition of the light-harvesting complexes (LHCs) and of a
number of other components of their photosynthetic machinery (for
review, see Falkowski and Laroche, 1991
; Anderson et al., 1995). On
shorter time scales, they react to an unbalance between light intensity
and photosynthetic capacity (e.g. due to a change in light intensity,
temperature, or nutrient supply) by rapid structural modifications
within the LHC of PSII (Horton et al., 1996; Bassi and Caffarri, 2000).
These modifications lead to a decrease in the fluorescence yield
designated as the non-photochemical quenching (NPQ) of chlorophyll
(Chl) a fluorescence. NPQ is supposed to dissipate excess
excitation energy through a harmless non-radiative pathway. The
partitioning of absorbed energy between transfer to the reaction center
and photoprotective non-radiative dissipation is controlled by the
trans-thylakoid pH gradient (for recent review, see Müller et
al., 2001) and by the reversible conversion of epoxidized to
de-epoxidized forms of xanthophylls (the so-called xanthophyll cycle;
Gilmore, 1997). The molecular mechanisms of photoprotection have mostly
been studied in higher plants (Demmig-Adams and Adams, 1996). Several
mutants of Arabidopsis and Chlamydomonas reinhardtii
with modified violaxanthin (the epoxidated xanthophyll cycle pigment
form) content and NPQ extent have been investigated and a specific role
has been attributed to the PsbS protein (Niyogi, 1999; Müller et
al., 2001). However, the field remains controversial due to the obvious
complexity of photoprotective mechanisms. In this sense, the use of
diatoms to study the role of xanthophylls in photoprotection may have important advantages.
Diatoms are well known to flourish in turbulent waters (Harris, 1986),
where the amount of light available to the phytoplanktonic unicellular
organisms is highly unpredictable. The deep vertical mixing
continuously sweeps them up and down, exposing the cells to very large
short-term changes in light intensity on a time scale of minutes to
hours. The organization of the photosynthetic apparatus in diatoms
differs in many respects from that of higher plants. The thylakoids are
loosely appressed and organized in extended bands of three, and the PSI
and PSII are not segregated in different domains (Pyszniak and Gibbs,
1992). The LHCs, which contain Chl a, Chl c,
fucoxanthin, and the xanthophyll cycle pigment diadinoxanthin (DD;
Brown, 1988), are equally distributed among appressed and nonappressed
regions (Pyszniak and Gibbs, 1992) and there is no evidence of any
state transitions (Owens, 1986). The LHCs subunits are made of several
highly homologous proteins encoded by a multigene family (fucoxanthin
Chl proteins; Bhaya and Grossman, 1993). The CP26 and CP29 subunits
present in higher plants are not found in diatoms (Müller et al.,
2001) and the existence of the PsbS protein has yet to be proven. The
supramolecular organization of PSII within the membrane remains
unknown. When the cells are suddenly exposed to high-light intensity,
an NPQ is rapidly developed (Demers et al., 1991; Ting and Owens, 1993; Arsalane et al., 1994; Olaizola and Yamamoto, 1994; Olaizola et al.,
1994; Casper-Lindley and Bjorkman, 1998). NPQ is associated with a
xanthophyll cycle (the DD cycle), which differs from that of higher
plants. The DD cycle converts the mono-epoxide carotenoid DD into the
de-epoxide form diatoxanthin (DT) under high light, and DT back into DD
under low light or in darkness (Arsalane et al., 1994; Casper-Lindley
and Bjorkman, 1998).
In diatoms, the DD content can be modulated by the light regime to
which culture is exposed (Willemoës and Monas, 1991; Arsalane et
al., 1994; Casper-Lindley and Bjorkman, 1998; Mouget et al., 1999). We
were able to define an intermittent light (IL) regime in which cells of
the diatom Phaeodactylum tricornutum had an increased DD
content but unchanged PSII antenna size. By comparing cells with either
a low or high DD content, we studied the influence of the DD pool size
on the DD cycle activity and NPQ formation under high light and
investigated its functional role in protecting PSII from
overexcitation. All results obtained demonstrate that a larger pool of
DD leads to a more effective photoprotection.
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RESULTS |
Comparison of the Photosynthetic Apparatus of Continuous Light (CL)
and IL Cells
P. tricornutum cells with different DD contents were
obtained from cultures grown under different illumination regimes. CL cultures were grown using a 16-h-light/8-h-dark cycle. IL cultures were
grown under a 5-min-light/55-min-dark cycle. In both cases, the light
intensity was 40 µE m
2
s1, which was weak enough not to induce any
de-epoxidation of DD. The growth rate under IL illumination was reduced
by a factor of 10, which is nearly accounted for by the 8-fold lower
total amount of light supplied.
The DD content of IL cells was 2 times higher than that of CL
cells (Table I). Otherwise, no
significant differences were found in their pigment contents. Cell size
and chloroplast ultrastructure were similar for the two types of cells.
It is important to note that the PSII antenna size was the same in the
two types of cells, in their dark-adapted state, as indicated by their
equal fluorescence yield rise time upon illumination in the presence of
DCMU (Table I). This was confirmed by the fact that equal flash
intensities were required for one-half saturation of the steady-state
O2 yield per flash with dark-adapted IL and CL
cells.
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Table I.
Pigment composition and PSII properties of the CL
and IL cells
Pigment contentsa and PSII properties of
P. tricornutum grown under CL and IL illumination are
compared. YSS is the steady-state O2 yield per
single-turnover flash, a measure of the concentration of active PSII
reaction centers.
Fv/Fm
(Fv = Fm Fo) is the quantum yield of excitation
trapping by PSII. The one-half rise time of the fluorescence induction
at a given light intensity in presence of
3-(3',4'-dichlorophenyl)-1,1-dimethylurea (DCMU) is a measure of the
PSII antenna size. I1/2 of YSS is the flash
intensity required for half-saturation of YSS, reciprocal
measure of the PSII antenna size. Isat is the minimum
intensity of continuous light that saturates the rate of O2
emission. Data (±SD) are the average of six measurements
for pigment contents and three measurements for other parameters. All
data except Isat refer to samples without DT and NPQ.
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NPQ Evaluated by Stern-Volmer Coefficients (SVo and
SVm) in CL and IL Cells
Figure 1 shows typical PAM
fluorometer recordings of the Chl fluorescence yield changes induced by
illumination of CL cells (A) and IL cells (B), as detected by a
nonactinic modulated beam. The cells were illuminated for 5 min,
indicated by up and down arrows, at an intensity of 450 µE
m2 s1 that was just
saturating for the rate of oxygen evolution. The spikes result from
temporary accumulation of QA
(reduced primary acceptor of PS II) by short pulses of much stronger illumination, applied at regular intervals to probe the fluorescence yield in the absence of photochemical quenching. The data are normalized to the maximum fluorescence yield,
Fm, reached when such a pulse is applied to
dark-adapted cells that show only photochemical quenching and have a
low fluorescence yield (Fo). During the
5-min illumination, a large NPQ rapidly developed, as shown by the
lower maximum yields reached by the high intensity pulses
(Fm'). This NPQ also affects the
fluorescence yield of "open" PSII, as seen by the lower minimum
yield (Fo'). We will use this value to
indicate Fo', but note that its real value
might be even somewhat lower because the presence of some remaining
QA at this point in time
cannot be ruled out. The minimum of Fo' variations is reached in less than 2 min. Some NPQ may already have
disappeared, but the corresponding Fm'
values show that NPQ relaxation is still small at this point. Recovery
of the dark-adapted state took about 15 min in CL cells (Fig. 1A). In
IL cells, NPQ induced by the same illumination was much more
pronounced, appeared more rapidly, and disappeared more slowly than in
CL cells (Fig. 1B).
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Figure 1.
Fluorescence yield recordings by the
pulse-amplitude-modulated (PAM) fluorometer of CL cells (A) and
IL cells (B). After a few minutes of darkness (with the modulated
detecting beam on), cells were illuminated for 5 min at 450 µE
m2 s1 (between up and
down arrows). At regular intervals, strong light pulses were fired to
probe the fluorescence yield without photochemical
quenching.
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The extent of NPQ is usually quantified as F/F' 1, which should be proportional to the quencher concentration if a
simple Stern-Volmer relation applies (see "Materials and Methods").
We will use the Stern-Volmer coefficients SVo and
SVm to denote
Fo/Fo' 1 and
Fm/Fm' 1, respectively. The induction kinetics of SVo and
SVm in the two types of cells is shown in Figure
2 for light intensities of 450 µE
m2 s1 (A, just
saturating light, Table I) and 2 mE m2
s1 (B, full sunlight, Long et al., 1994). The
white and black symbols refer to CL and IL cells, respectively, and
circles and triangles refer to SVo and
SVm, respectively. In all conditions, the
kinetics were similar: The major fast phase already visible in Figure 1 was followed by a much slower increase. Figure
3 shows SVo and SVm induced by 5 min of illumination as a
function of light intensity. CL cells required a somewhat higher light
intensity for NPQ induction than IL cells, but for both a large part of
the maximum NPQ was already induced at a light intensity of 0.5 mE
m2 s1 that was just
saturating for oxygen evolution.
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Figure 2.
NPQ as a function of illumination duration for two
light intensities: 450 µE m2
s1 (A) and 2 mE m2
s1(B). Quenching of
Fm (circles, SVm) and
of Fo (triangles,
SVo) in CL cells (white symbols) and IL cells
(black symbols). The data were obtained from measurements as in Figure
1.
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Figure 3.
NPQ as a function of light intensity for a fixed
illumination duration of 5 min. SVm (circles) and
SVo (triangles) for CL cells (white symbols) and
IL cells (black symbols).
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Correlation of NPQ and DT Accumulation
The kinetics of DT accumulation was similar to that of NPQ
formation. No DT could be detected in dark-adapted cells. The amount of
DT formed by de-epoxidation of DD results from the competition between
epoxidation and de-epoxidation (activated by the proton gradient). At
the onset of light, the first phase corresponds to the rapid
transformation of DD to DT. The equilibrium reached at the end of the
5-min illumination depends on the light intensity: At low light,
epoxidation competes efficiently with de-epoxidation, then
de-epoxidation becomes more efficient with increasing light intensities
and a quasi-steady state is reached when the equilibrium is strongly
shifted in favor of DT.
The maximum amount of DT formed, measured after 1 h of
illumination at 2 mE m2
s1, was 5 and 12.5 mol of DT (100 mol of Chl
a)1 for CL and IL cells,
respectively, corresponding to the de-epoxidation of 50% and 65% of
the total DD pool. As shown in Figure 4,
SVo and SVm were found to
be proportional to the DT concentration for both types of cells.
Although the regression coefficients for SVm were
very similar (0.95 for CL and 1.04 for IL cells), those for
SVo were quite different for the two types of
cells: 0.39 for CL and 0.72 for IL cells. The ratio of
SVo/SVm would have been
equal to Fo/Fm
(near 0.22 in both cell types) if the photochemical and
non-photochemical quenchers were randomly competing for excitations in
the same homogeneous pigment bed. The ratio SVo/SVm is, however, 0.41 for CL cells and 0.69 for IL cells. If the values used for
Fo' were slightly overestimated, even
larger values of SVo would result after
correction.
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Figure 4.
Correlation of NPQ and DT concentration.
SVm (circles) and SVo
(triangles) for CL cells (white symbols, shown enlarged in the inset)
and IL cells (black symbols). Linear regressions were:
SVm(CL) = 0.95 [DT],
SVm(IL) = 1.04 [DT],
SVo(CL) = 0.39 [DT], and
SVo(IL) = 0.72 [DT] with 0.98 < R2 <0.99. [DT]/[Chl a] was
determined by HPLC on cells sampled at the end of the light treatment.
The duration of light was varied from 30 s to 60 min and the light
intensity from 0.05 to 2 mE m2
s1.
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Decrease of the Quantum Efficiency of PSII by NPQ
Because NPQ dissipates part of the excitation energy, it is
expected to decrease the quantum efficiency of PSII. This was verified
by measurements of the amount of oxygen produced in short flashes as a
function of flash energy (Fig. 5). The
saturation curves of dark-adapted CL and IL cells (white symbols) were
almost identical, confirming that the PSII antenna size was the same in
both culture conditions. The flash illumination used, 20 single-turnover flashes in 10 s, corresponds to a very low
continuous intensity and did not induce any NPQ. After exposure to
high-light intensity, the saturation curves for flash-induced oxygen
evolution were shifted to higher flash energies (black symbols). These
measurements were taken 7 min after the light treatment (the time
required for sedimentation on the electrode) and were compared with the fluorescence yield measured at the same time after illumination, which
is already substantially less quenched by NPQ than 2 min after
illumination (see Fig. 1). The remaining quenching of
Fo at this time was equal to the
decrease of the quantum yield of O2
evolution, as indicated by the reciprocal of the one-half-saturating flash intensity (inset, Fig. 5). This result is consistent with Equation 3 in "Materials and Methods" and may indicate that PSI fluorescence does not significantly contribute to
Fo or is similarly quenched by NPQ.
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Figure 5.
Steady-state O2 yield per
flash (YSS) as a function of flash intensity.
Dark-adapted (SVo = 0) CL cells (white circles)
and IL cells (white triangles), CL cells at SVo = 0.5 (black circles) and IL cells at SVo = 1 (black triangles). SVo was measured
simultaneously to YSS (after 7 min of darkness)
with the PAM fluorometer. The intermediary curve correspond to CL cells
with a maximal SVo value of 1.5 after cessation
of illumination (15 min at 2,000 µE m2
s1) lowered to 0.5 min when the flash series
was fired 7 min later. The curve to the right corresponded to IL cells
with a maximal SVo value of 4 after cessation of
illumination (5 min at 450 µE m2
s1) lowered to 1, when the flash sequence was
fired. Inset, Relative reciprocal light intensity needed for one-half
saturation of YSS (I1/2 [DT = 0]/I1/2) versus
Fo'/Fo.
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If NPQ decreases the quantum yields of fluorescence and
O2 evolution equally also at stronger quenching,
when the available flash energies were insufficient to determine the
one-half-saturating intensity, the energy dissipation measured by
1 Fo'/Fo equals the down-regulation of PSII by NPQ. Figure
6 shows the development of the energy
dissipation in the antenna of open PSII centers for the two types of
cells as a function of illumination time at 0.45 and at 2 mE
m2 s1. The energy
dissipation reached values up to 90% and was much higher for IL cells
than for CL cells at both light intensities, especially after short
illumination times (enlarged in the inset).
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Figure 6.
Energy dissipation in the antenna of open PSII
reaction centers (1 Fo'/Fo) as a
function of illumination duration. CL cells (white symbols) and IL
cells (black symbols) at 450 µE m2
s1 (dashed lines) and 2 mE
m2 s1 (black lines).
Inset, First 5 min enlarged.
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Protection by NPQ against Photoinhibition
Because CL and IL cells appeared to differ only in the capacity of
energy dissipation by DT, their comparison provides a unique opportunity to verify the functional role of this process in protecting PSII against photoinhibition. Photoinhibition was evaluated by monitoring O2 evolution at the Clark electrode
and measuring the time needed to reach the compensation point (Arsalane
et al., 1994). As predicted by Arsalane et al. (1994), the larger the DD content, the longer time is needed to reach the compensation point
(not shown).
An additional method for evaluation of photoinhibition was used. Figure
7 shows the decrease of PSII activity
measured as the steady-state O2 yield per flash
(YSS) after illumination at 0.45 or 2 mE
m2 s1 followed by 7 min
of dark adaptation. Photoinhibition kinetics were faster for CL cells
(white symbols) than for IL cells (black symbols), especially during
the first 5 min. After 1 h of illumination, the remaining
percentage of active PSII centers was lower for CL cells under the two
light intensities. The difference between IL and CL cells is actually
larger than Figure 7 suggests, because especially for IL cells the
recovery from NPQ is much slower than 7 min (Fig. 1) and saturation of
YSS could not be reached with the available flash
energy. Because 85% or 90% of the energy was dissipated by NPQ after
1 h at 0.45 or 2 mE m2
s1, respectively (Fig. 6), the flash saturation
curve for dark-adapted cells in Figure 5 indicates that only 87% or
79% saturation of YSS could be reached. The
exact correction has not been done because combining the information
from the different data in Figures 5 through 7 in this way seems too
daring, but the corrections may obviously be large for IL cells. The
correction would leave essentially no evidence for photoinhibition of
IL cells at 0.45 mE at all. For CL cells, no such correction applies
because NPQ largely disappeared in 7 min and even at 65% energy
dissipation (Fig. 6), the flashes would still be more than 95%
saturating (Fig. 5). The fast phase of photoinhibition seen during the
first few minutes at high-light intensity correlates with the time
required for accumulation of DT and NPQ and illustrates the importance
of the photoprotective effect. Such a short delay in the buildup of the
protection is unlikely to be of consequence even in the turbulent
native environment of P. tricornutum.
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Figure 7.
Photoinhibition kinetics of PSII as a function of
illumination duration. CL cells (white symbols) and IL cells (black
symbols) at 450 µE m2
s1 (squares) and 2 mE
m2 s1 (circles). After
the light treatment, cells were allowed to settle on a rate electrode
for 7 min in darkness. PSII activity was estimated by measurement of
the steady-state O2 yield per flash after a
sequence of twenty single-turnover flashes.
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DISCUSSION |
DD Enrichment and the Light-Induced DT Accumulation
P. tricornutum cultures in stationary phase show an
inverse relation between growth rate and DD content (Arsalane et al., 1994). To slow down the growth rate and to keep the cells in the exponential phase of growth, we grew P. tricornutum under an
IL regime. When P. tricornutum are grown under such a
regime, the Chl and fucoxanthin content per cell do not change, whereas
the size of the DD pool is increased, in striking contrast to the effect of intermittent illumination on higher plants, which leads to a
drastic reduction of PSII antenna size. IL-grown peas
(Pisum sativum) are devoid of all LHCII proteins,
with the exception of CP26, one of the minor inner antenna Chl-binding
proteins (Jahns and Junge, 1992). This way, a specific increase in DD
content could be induced without significant change in the amounts of other pigments, PSI/PSII ratio, or PSII antenna size and quantum yield.
Upon exposure to high-light intensity, a substantial de-epoxidation of
DD occurred, largely within a few minutes. This in agreement with DD
cycle activity previously reported (Arsalane et al., 1994; Olaizola et
al., 1994; Casper-Lindley and Bjorkman, 1998). The larger pool of DD in
IL cells, and its more extensive de-epoxidation, led to much larger
accumulation of DT during high-light illumination. The maximal DT
accumulation is 5 mol of DT (100 mol of Chl
a)1 in CL cells and 12.5 mol of DT
(100 mol of Chl a)1 in IL cells. In
higher plants, Ruban et al. (1999) have estimated that each LHCII
monomer can bind at least one violaxanthin. Upon activation of the
violaxanthin de-epoxidase, the highest de-epoxidation state was found
for the main LHCII components and the lowest for CP29. Very little is
known about the nature of the subunits of LHC in diatoms (Bhaya and
Grossman, 1993) and it seems premature to speculate on their DT-binding
sites at this stage.
Correlation of Light-Induced DT Accumulation and NPQ
The Stern-Volmer parameters SVo and
SVm relate the NPQ on
Fo and Fm to
the quencher concentration (Bilger and Björkman, 1990; Gilmore
and Yamamoto, 1991; Olaizola and Yamamoto, 1994). Random quenching in a
homogenous antenna would produce a normalized fluorescence yield as
depicted by Equation 1 (see "Materials and Methods"). For the
quenching of Fm, DT is the only quencher
present in the LHC of PSII, the photochemical quenching being totally
suppressed. In that case, the slope of SVm is the
same for IL and CL cells. (SVm)IL can reach values
higher than (SVm)CL, the
difference being only due to the difference in the maximal
concentration of DT that can be formed (see Fig. 4). For the
Fo level, the photochemical quencher (Q) is
present (PSII centers are open) and the two types of quenchers compete
for the trapping of the excitation. If the quenchers were randomly
competing in a homogeneous antenna, the slope of
SVo should be
Fo/Fm × slope
of SVm (see Eq. 2 in "Materials and
Methods"). It is not the case: The observed
SVo/SVm ratios were much
larger than
Fo/Fm,
especially for IL cells. The discrepancy is far too large to be
substantially affected by possible errors in the determination of
the extent of photochemical quenching due to quenching by the
plastoquinone pool or incomplete oxidation of
QA after illumination. Our
results provide an additional indirect proof that excitation transfer
to the open PSII reaction centers becomes rate limiting for trapping by
open PSII reaction centers, causing a heterogeneity of the Chl
fluorescence yield in the PSII antenna. The selective increase of
SVo indicates that the Chls involved contribute a
larger fraction of the emission at Fo than at Fm.
Functional Significance of Energy Dissipation by DT and
Photoinhibition
Most studies on NPQ consider only the quenching of
Fm. Although the quenching of
Fm presumably helps to diminish singlet
oxygen production in the antenna by reducing the yield of Chl triplet formation, the quantitative significance of this protective effect under physiological conditions is probably negligible. The triplet yield of Chl a is twice the fluorescence yield and in an
aerobic environment its very fast transfer to carotenoids followed by intersystem crossing to the ground state is essential anytime. Only the
reaction center Chl itself cannot be protected this way (van Gorkom and
Schelvis, 1993), but at Fm the PSII
reaction center does not quench fluorescence (Thielen and van Gorkom,
1981). Hence, the probability that the triplet state is formed there,
rather than in the antenna, is as small as the fraction of Chl
a contained in the reaction center. At
Fo, on the other hand, most excitations cause a charge separation. If that cannot be stabilized by secondary electron transfer reactions due to saturation of electron transport, it
will most likely recombine to the reaction center triplet state (van
Gorkom, 1985, 1986). Any mechanism to avoid that would be physiologically significant and could bring a decisive advantage under
continually and widely changing light intensity. Our results show that
the xanthophyll cycle in P. tricornutum provides such a mechanism.
The quenching of Fo is a direct measure of
the extent of down-regulation of PSII electron transport resulting from
NPQ. We have shown that energy dissipation by DT equally decreases the quantum yield of O2 evolution and the quantum
yield of fluorescence emission when the PSII centers are open. The
corresponding effect in higher plants may be small (see Santabarbara et
al., 2001), but our results clearly confirm the large decrease of PSII
quantum efficiency by NPQ in P. tricornutum, reported by
Koblizek et al. (2001). Moreover, this unusual capacity for
down-regulation of PSII is adjusted to the illumination conditions
during growth.
Finally, we note an intriguing observation: After 1 h at 2 mE
m2 s1, CL cells lost up
to 70% of their O2-evolving PSII reaction
centers, and yet no deviation from the proportionality between
SVo, SVm, and DT is seen in
the inset of Figure 4. Apparently, there is nothing special about the
fluorescence yield, including the variable fluorescence, of this
"inactivated PSII." This indicates that these diatoms are not
photoinhibited in the classical sense and, in fact, Olaizola et al.
(1994) already mentioned the absence of D1 photodamage in P. tricornutum. Evidence for the induction of an electron transport
cycle around PSII that competes with oxygen evolution at high-light
intensity will be presented elsewhere (Lavaud et al., 2002).
The phenomena reported here are probably not restricted to P. tricornutum. Other marine planktonic diatoms like
Chaetoceros muelleri (Olaizola et al., 1994),
Thalassiosira pseudonana (Demers et al., 1991),
Haslea ostrearia (Mouget et al., 1999),
and freshwater planktonic diatoms like Nitzschia
palea (Willemoës and Monas, 1991) are able to
synthesize large amounts of de-epoxidizable xanthophylls. It seems
likely that the large and adjustable capacity of DT accumulation, and
the fast and efficient photoprotection by the associated NPQ, are
general properties of planktonic diatoms and play an important role in
their successful adaptation to the strongly fluctuating light intensity
in their natural habitat (Harris, 1986).
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MATERIALS AND METHODS |
Cultivation and Preparation of Cells
Phaeodactylum tricornutum Böhlin cells were
grown photoautotrophically in sterile natural seawater F/2 medium
(Guillard and Ryther, 1962). Cultures of 300 mL were incubated at
18°C in airlifts continuously flushed with sterile air. They were
illuminated at a light intensity of 40 µE m2
s1 with white fluorescent tubes (Claude, Blanc Industrie,
France) with a 16-h-light/8-h-dark cycle for CL cells or with
5-min-light/55-min-dark cycle for IL cells. Cells were harvested during
the exponential phase of growth, centrifuged at 3,000g
for 10 min, and resuspended in their culture medium to a final Chl
a concentration of 10 µg Chl a
mL1. The algae were continuously stirred at 18°C under
low CL for CL cells and under IL for IL cells until measurement.
Pigment Contents
Pigment analyses were performed by HPLC as previously described
(Arsalane et al., 1994). Cells collected from the PAM fluorometer vial
(see below) were frozen in liquid nitrogen. Pigments were extracted
with a methanol:acetone (70:30 [v/v]) solution. The extinction
coefficients used were the same as in Berkaloff et al. (1990) for Chl
and as Johansen et al. (1974) for DD and DT. Cell counts were performed
with a Thoma hematocymeter, using the public domain NIH Image program
(National Institutes of Health, Rockville, MD).
PSI Reaction Center (P700) Concentration
P700 was determined as described by Newman and
Sherman (1978). Cells were first adjusted to the same Chl
a concentration and then broken by sonication and kept
on ice. The light-induced absorption decrease at 700 nm was measured
with a DW-2 Aminco spectrophotometer (American Instrument Co., Olis
Incorporated, Bogart, GA) in dual-beam mode (reference at 730 nm) in the presence of 6 mM sodium ascorbate and 300 to 600 µM methylviologen; 30 µM DCMU was added to
prevent P700+ reduction by linear electron
flow. Stock solutions of sodium ascorbate (0.2 M in
distilled water, Sigma, St. Louis), methylviologen (Sigma, 10 mM in distilled water), and DCMU (Sigma, 10 mM
in absolute ethanol) were freshly prepared.
PSII Antenna Size Assessed by Fluorescence Emission Induction in
the Presence of DCMU
PSII Chl fluorescence induction kinetics of dark-adapted cells
were measured at room temperature in a laboratory-built continuous fluorometer. The setup and the data acquisition were described previously by Ritz et al. (1999). Samples were adjusted to a final concentration of 4 µg Chl a mL1
and were dark adapted for 20 min before the experiment. Twenty micromolar DCMU was added 5 min before the end of the dark period.
O2 Concentration and Photosynthetic Light Response
Curves
O2 concentration was measured with a DW1-Clark
electrode (Hansatech Ltd., King's Lynn, UK) at 18°C. White
light of adjustable intensity (measured with a PAR-sensor, LI-COR,
Lincoln, NE) was provided by KL-1500 quartz iodine lamp (Schott,
Mainz, Germany). Cell culture samples were dark adapted for 20 min
before measurement. Photosynthetic light-response curves were obtained
by illuminating a 2-mL sample during 5 min at various intensities. A
new sample was used for each intensity.
Chl Fluorescence Yield Measurements
The Clark electrode set-up was modified to allow simultaneous
measurement of oxygen concentration and Chl fluorescence by a PAM-101
fluorometer (Walz, Effeltrich, Germany) as described previously (Ritz
et al., 1999). The fluorescence excited by a very weak (nonactinic)
modulated 650 nm of light was measured. After 20 min of dark
adaptation, continuous actinic light of adjustable intensity was
switched on. Six hundred-millisecond pulses of white light (4 mE
m2 s1) were admitted by an electronic
shutter (Uniblitz, Vincent, Rochester, NY, opening time 2 ms)
placed in front of a KL-1500 quartz iodine lamp continuously switched
on to monitor the NPQ evolution. The average fluorescence (acquisition
time 33 µs) measured during the last 400 ms of the pulse was taken as
Fm or Fm'. Data
were recorded with a microcomputer through a 12-bit analog-digital interface and the system was driven by homemade software (Arsalane et
al., 1993). For each experiment, 2 mL of cell suspension was used.
Sodium bicarbonate (Labosi, Elancourt, France) was added at a
concentration of 4 mM from a freshly prepared 0.2 M stock solution in distilled water to prevent any
limitation of the photosynthetic rate by carbon supply.
Standard fluorescence nomenclature was used (van Kooten and Snel,
1990). Fo and Fm
are defined as the fluorescence yield of dark-adapted cells and the
maximum fluorescence reached in such cells during a saturating pulse of
white light, respectively. The quantum yield of excitation trapping by
PSII is the ratio Fv/Fm, where
Fv is the variable part of the fluorescence
emission and is equal to Fm Fo. NPQ is quantified as
F/F' 1, where F' is the
fluorescence yield in the presence of quenching (Bilger and
Björkman, 1990; Gilmore and Yamamoto, 1991). We use the
Stern-Volmer parameters SVo and SVm to indicate
NPQ of Fo and NPQ of
Fm, respectively. The fluorescence yields
Fo, Fm,
Fo', and Fm' are
indicated in Figure 1A.
The following formalism was used to relate the Stern-Volmer equations
with the amounts of photochemical (Q for quinones) and non-photochemical (DT) quenchers. The decay rate of the excited state
of Chl a molecules in PSII is:
where kF is the rate
sum of the constant of fluorescence emission,
kIC is the sum of the rate constant of
intersystem crossing to the Chl a triplet state and the
rate constant of internal conversion to the ground state,
kQ [Q] is the rate constant of
photochemical quenching, and kDT [DT] is the rate constant of NPQ.
The fluorescence yield is:
In the absence of NPQ, Q is the only quencher and
when Q = 0 and when Q is at its maximal concentration
then,
In the presence of the quencher DT, if Q = 0,
![<FR><NU>F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>m′</UP></SUB></DE></FR>=1+k′<SUB>DT</SUB>[DT] <UP>and SV<SUB>m</SUB></UP>=<FR><NU>F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>m′</UP></SUB></DE></FR>−1=k′<SUB>DT</SUB>[DT]](/content/vol129/issue3/fulltext/1398/img006.gif)
|
(1)
|
In the presence of both quenchers,
Then,
![<UP>SV</UP><SUB><UP>o</UP></SUB>=<FR><NU><IT>F</IT><SUB><UP>o</UP></SUB></NU><DE><IT>F</IT><SUB><UP>o′</UP></SUB></DE></FR>−1=<FR><NU><IT>F</IT><SUB><UP>o</UP></SUB></NU><DE>F<SUB><UP>m</UP></SUB></DE></FR>k′<SUB><IT>DT</IT></SUB>[<IT>DT</IT>]=<FR><NU><IT>F</IT><SUB><UP>o</UP></SUB></NU><DE><IT>F</IT><SUB><UP>m</UP></SUB></DE></FR><UP>SV<SUB>m</SUB></UP>](/content/vol129/issue3/fulltext/1398/img008.gif)
|
(2)
|
O2 Yield per Flash
The relative O2 yield produced per flash during a
sequence of single-turnover saturating flashes at a frequency of 2 Hz
was measured with a rate electrode described by (Lemasson and Etienne, 1975). Short (5-µs) saturating flashes were produced by a Strobotac (General Radio Co., Concord, MA). O2 evolution was
monitored at 18°C. For the control sequences, cells were first dark
adapted during 20 min and then deposited on the electrode. Both control and illuminated samples taken from the PAM fluorometer were allowed to
settle on the electrode for 7 min in darkness before measurement. The
steady-state O2 yield per flash (YSS) was taken
to be the average yield of the last four flashes of a series of 20 flashes when the classical four-step oscillations due to the S states cycle (Kok et al., 1970) was almost damped. YSS was used to
assess the relative concentration of O2-evolving PSII
reaction centers. For the saturation curves of YSS, the
intensity of the flashes was varied with neutral density filters.
Samples were taken from the PAM fluorometer after illumination and
deposited on the electrode. For each flash intensity, a new sample was
used. In parallel, fluorescence yield was measured with the PAM setup
at the same time after illumination.
The reciprocal of the one-half-saturating flash intensity as a measure
of the antenna size was expressed as follows. The quantum yield of
photochemistry is given by the equation:
and should be inversely proportional to the amount of
light needed to activate a given fraction of the reaction centers, so that:
![<FR><NU>I<SUB>1/2</SUB></NU><DE>I<SUB>1/2[<UP>DT</UP>=0]</SUB></DE></FR>=<FR><NU>&PHgr;<SUB><UP>Q</UP>[<UP>DT</UP>=0]</SUB></NU><DE>&PHgr;<SUB><UP>Q</UP></SUB></DE></FR>=<FR><NU>k<SUB>F</SUB>+k<SUB>IC</SUB>+k<SUB>Q</SUB>[<UP>Q</UP>]+k<SUB>DT</SUB>[DT]</NU><DE>k<SUB>F</SUB>+k<SUB>IC</SUB>+k<SUB>Q</SUB>[<UP>Q</UP>]</DE></FR>=<FR><NU>F<SUB><UP>o</UP></SUB></NU><DE>F<SUB><UP>o′</UP></SUB></DE></FR>](/content/vol129/issue3/fulltext/1398/img010.gif)
|
(3)
|
with I1/2 the light intensity needed for
one-half saturation of YSS in the presence of DT and
I1/2 [DT = 0] the light intensity needed for
one-half saturation of YSS in absence of DT.
We thank Prof. Jean-Claude Duval for experimental
assistance and helpful discussions, Christiane Lichtlé for
electron microscopic photographs, and Dr. Krishna Niyogi and Andrew
Pascal for critical reading of the manuscript.
Received December 21, 2001; returned for revision February 21, 2002; accepted March 18, 2002.
Article, publication date, and citation information can be found at
www.plantphysiol.org/cgi/doi/10.1104/pp.002014.
TOP
ABSTRACT
INTRODUCTION![<FR><NU>d[*]</NU><DE>dt</DE></FR>=<LIM><OP>∑</OP></LIM>k[*] <UP>with </UP><LIM><OP>∑</OP></LIM>k=k<SUB>F</SUB>+k<SUB>IC</SUB>+k<SUB>Q</SUB>[<UP>Q</UP>]+k<SUB><IT>DT</IT></SUB>[<UP>DT</UP>]](/content/vol129/issue3/fulltext/1398/img001.gif)
![F<SUB><UP>o</UP></SUB>=<FR><NU>k<SUB>F</SUB></NU><DE>k<SUB>F</SUB>+k<SUB>IC</SUB>+k<SUB>Q</SUB>[<UP>Q</UP>]</DE></FR>](/content/vol129/issue3/fulltext/1398/img004.gif)
![<FR><NU>F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>o</UP></SUB></DE></FR>=1+<FR><NU>k<SUB>Q</SUB></NU><DE>k<SUB>F</SUB>+k<SUB>IC</SUB></DE></FR>[<UP>Q</UP>]=1+k′<SUB>Q</SUB>[<UP>Q</UP>]](/content/vol129/issue3/fulltext/1398/img005.gif)
![<FR><NU>F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>o′</UP></SUB></DE></FR>=1+k′<SUB>Q</SUB>[Q]+k′<SUB>DT</SUB>[DT]=<FR><NU>F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>o</UP></SUB></DE></FR>+k′<SUB>DT</SUB> <UP>and </UP><FR><NU>F<SUB><UP>o</UP></SUB></NU><DE>F<SUB><UP>o′</UP></SUB></DE></FR>=<FR><NU>F<SUB><UP>o</UP></SUB>×F<SUB><UP>m</UP></SUB></NU><DE>F<SUB><UP>m</UP></SUB>×F<SUB><UP>o′</UP></SUB></DE></FR>=1+<FR><NU>F<SUB><UP>o</UP></SUB></NU><DE>F<SUB><UP>m</UP></SUB></DE></FR>k′<SUB>DT</SUB>[DT]](/content/vol129/issue3/fulltext/1398/img007.gif)
![&PHgr;<SUB><UP>Q</UP></SUB>=<FR><NU>k<SUB>Q</SUB>[<UP>Q</UP>]</NU><DE>k<SUB>F</SUB>+k<SUB>IC</SUB>+k<SUB>Q</SUB>[<UP>Q</UP>]+k<SUB>DT</SUB>[DT]</DE></FR>](/content/vol129/issue3/fulltext/1398/img009.gif)
pH-dependent fluorescence quenching and its photoprotective role in the unicellular red alga Rhodella violacea.
Photosynthetica
37: 267-280
