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Plant Physiol. (1998) 116: 1551-1562
Photosystem II Cyclic Heterogeneity and Photoactivation in the
Diazotrophic, Unicellular Cyanobacterium
Cyanothece
Species ATCC 511421
Pascal C. Meunier,
Milagros S. Colón-López, and
Louis
A. Sherman*
Department of Biological Sciences, Purdue University, West
Lafayette, Indiana 47907
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ABSTRACT |
The unicellular, diazotrophic
cyanobacterium Cyanothece sp. ATCC 51142 demonstrated
important modifications to photosystem II (PSII) centers when grown
under light/dark N2-fixing conditions. The properties of
PSII were studied throughout the diurnal cycle using
O2-flash-yield and pulse-amplitude-modulated fluorescence techniques. Nonphotochemical quenching (qN)
of PSII increased during N2 fixation and persisted after
treatments known to induce transitions to state 1. The
qN was high in cells grown in the dark, and
then disappeared progressively during the first 4 h of light
growth. The photoactivation probability,  , demonstrated interesting
oscillations, with peaks near 3 h of darkness and 4 and 10 h
of light. Experiments and calculations of the S-state distribution
indicated that PSII displays a high level of heterogeneity, especially
as the cells prepare for N2 fixation. We conclude that the
oxidizing side of PSII is strongly affected during the period before
and after the peak of nitrogenase activity; changes include a lowered
capacity for O2 evolution, altered dark stability of PSII
centers, and substantial changes in qN.
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INTRODUCTION |
Many species of bacteria and cyanobacteria are capable of
N2 fixation. Nitrogenase catalyzes the
energetically expensive reduction of N2 to
NH3 and is rapidly and irreversibly inhibited
when extracted from cells in the presence of free
O2 (Fay and Cox, 1967 ; Haystead et al., 1970).
The N2-fixation trait is particularly interesting in cyanobacteria because of the apparent incompatibility of an O2-sensitive nitrogenase complex in a prokaryotic
organism capable of splitting water to produce
O2. Cyanobacteria have developed elegant
strategies to protect nitrogenase from inactivation by O2, including spatial separation, temporal
separation, and the induction of enzyme systems to destroy reactive
O2 byproducts (Fay, 1992). The strategies devised
by nonheterocystous, unicellular cyanobacteria that permit
N2 fixation and O2
evolution in the same cell are quite sophisticated (Bergman et al.,
1997). A number of these unicellular, diazotrophic species fix
N2 and evolve O2 at
different times during the diurnal cycle. Such temporal separation has
been seen in strains classified as Gloeothece sp.
(Mullineaux et al., 1981) and Synechococcus sp. (Mitsui et
al., 1986; Grobbelaar et al., 1987).
Synechococcus Miami BG 43511 and BG 43522 (Mitsui et al.,
1987) can be synchronized by growth under light/dark cycles so that they continue to cycle nitrogenase even during continuous light. Under
these conditions, O2 evolution and nitrogenase
cycled in a reciprocal way such that O2 evolution
was low when nitrogenase activity was high. A similar reciprocal
relationship between the two metabolic processes has been demonstrated
in Synechococcus RF-1 (Grobbelaar et al., 1987),
in which nitrogenase activity was found to be highest in the dark and
O2 evolution highest in the light. However, in
that study little was done to analyze the specific changes that
occurred during photosynthesis.
We have begun to study diurnal metabolic rhythms and their control in
the unicellular, diazotrophic cyanobacterium Cyanothece sp.
ATCC 51142. It has been demonstrated previously that this strain
possesses extraordinary diurnal metabolic periodicities that resemble
circadian rhythms (Reddy et al., 1993; Schneegurt et al., 1994). When
cultures were grown in the absence of fixed N2,
either under 12-h light/12-h dark cycles or continuous light, the cells
fixed N2 with peaks of activity approximately
every 24 h. The peak of N2 fixation activity
was very narrow, with a half-width of about 2.5 h, reaching a
maximum about D4 (Schneegurt et al., 1994). The burst of
N2 fixation was accompanied by intense respiratory activity (Colón-López et al., 1997; Meunier et
al., 1997).
The nonheterocystous, filamentous, diazotrophic cyanobacterium
Plectonema boryanum also shows metabolic rhythms under
N2-fixing conditions, and its photosynthetic
behavior has been studied (Misra and Desai, 1993; Misra and Tuli,
1994). Photosynthetic activity measurements and thermoluminescence and
77 K fluorescence spectra suggested that PSI activity increased and
PSII activity decreased during the period of N2
fixation, suggesting that PSII was specifically regulated and that
electron transport under diazotrophic growth conditions was regulated
by the redox state of the secondary quinone acceptor,
QB. Furthermore, it was proposed that the PSII
QB was involved in PSII down-regulation. However,
none of this previous work analyzed the phenomenon at the level of the
water-oxidation mechanism (the S-state cycle).
We have determined that the physiology of Cyanothece sp.
ATCC 51142 involves important changes in PSII reaction center
oligomerization, phycobilisome attachment, gene transcription, protein
accumulation, and activity throughout the 24-h cycles (Meunier et al.,
1997; M.S. Colón-López and L.A. Sherman, unpublished
observations). However, the fate of the S-state cycle and
Mn2+ and O2 production
under flashing light throughout these structural changes has not yet
been investigated. PSII is a logical target for down-regulation during
periods of N2 fixation, since
O2 is produced by the PSII centers and
nitrogenase is sensitive to O2. Accordingly, we
found that PSI, but not PSII, contributes to N2 fixation if the cultures are illuminated during the normally dark N2-fixation period in Cyanothece sp.
ATCC 51142 (Meunier et al., 1997). In the current study, we analyzed
O2-flash-yield experiments and
qN data and related them to our previous
results to understand PSII function under these conditions. Our results
suggest that the properties of the oxidizing side of PSII are strongly
modified by variable dark-inactivation processes and photoactivation
that can be extremely rapid.
Photoactivation is the process by which Mn2+
becomes bound to the PSII reaction center and forms into a structure
that is active in the oxidation of water and in utilizing charge
separations as an energy source (Cheniae and Martin, 1972; Engels et
al., 1994; Gleiter et al., 1995; Burnap et al., 1996). Each step
between PSII charge separations is identified by an S-state, and
different S-states are denoted by Sx, where
"x" ranges from 0 to 4. Upon reaching the state
S4, O2 is released and the
process starts again from the S0 state (Kok et
al., 1970). The concept of S-states has been expanded to include the
steps required to oxidize Mn2+ up to the
S0 state. These steps are also known as the
super-reduced S-states S 1,
S2, and S3 (Bader et
al., 1983; Kretschmann and Witt, 1993; Messinger and Renger, 1993;
Messinger et al., 1997).
The reduction of the Mn2+ structure produces
deactivations of S-states and can result in the loss of
O2-evolving activity and the loss of
Mn2+ (Cheniae and Martin, 1972; Burnap et al.,
1996). Deactivations result in a loss of PSII efficiency and a
randomization of PSII centers among S-states. The susceptibility to
spontaneous deactivations is limited to the S3
and S2 states in most wild-type organisms; however, in some cyanobacterial PSII mutants, as well as in
Cyanothece sp. (see ``Results''), deactivations lead to
the loss of O2-evolving activity. A mechanistic
model of the S-state mechanism was recently proposed that explicitly
incorporates photoactivation and deactivation probabilities (Meunier et
al., 1996).
We will provide experimental evidence, using
O2-flash yields, that the water-splitting
mechanism is subjected to severe disturbances during the adaptive
changes of the photosynthetic apparatus and cycling physiology in
Cyanothece sp. ATCC 51142. We postulate that these
disturbances are common in diazotrophic cyanobacteria that are
unicellular or filamentous but nonheterocystous, and that the data
presented here are representative of a class of PSII centers optimized
for functioning under harsher conditions than those normally seen in
higher plants. Some of this research was presented in preliminary form
at the Tenth International Photosynthesis Congress (Meunier et al.,
1995b).
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MATERIALS AND METHODS |
Growth Conditions and Chlorophyll Determination
Cyanothece sp. ATCC 51142 (formerly strain BH68) was
grown as previously described in ASP2 medium without
NaNO3, with shaking at 100 or 125 rpm, at 30°C,
under cool-white fluorescent illumination of approximately 50 µE
m2 s1 (Reddy et al.,
1993). Continuous-light-grown, stationary-phase cultures were
subcultured by dilution to a concentration of 106
cells mL1. Duplicate flasks were subcultured
12 h apart using the same stock culture to permit 24-h experiments
to be performed in 12 h. After dilution, each flask was
illuminated for 12 h. This protocol, the determination of cell
number, and nitrogenase measurements were as described previously
(Schneegurt et al., 1994). The chlorophyll concentration was determined
using the freeware "Chlorophyll" (available for downloading at
http://bilbo.bio.purdue.edu/~pmeunier/download.html), according to the cyanobacterial method of Arnon et al. (1974). For measuring the chlorophyll concentration in whole cells, a spectrophotometer (model DU-7, Beckman) was modified by adding cellophane tape (no. 810, Scotch 3M Magic Tape) to the light path before and after the cuvette to correct for light scattering by the
cells. This yielded a fast and reproducible assay that was linear from
0.01 absorbance unit up to about 0.5 absorbance unit, and that was
within 10% of chlorophyll determinations performed using 80% acetone
extraction.
O2 Evolution/Flash-Yield Experiments
O2 yields after light flashes were measured
using a bare Pt electrode onto which cells were deposited by
centrifugation. Signals were amplified using an electronic circuit
based upon the design of Meunier and Popovic (1988). Cells (40 µg of
chlorophyll) were harvested by centrifugation at 10,000g in
a rotor (model JA-20, Sorvall) for 10 min, and resuspended in 200 µL
of buffer of the same osmolarity as the culture medium, but containing
only 340 mm NaCl and 10 mm Hepes, pH 7.5. The
200 µL was then centrifuged for 5 min at 1,000 rpm on the electrode
in a swing-out rotor (model HB4-A, Sorvall). The slow centrifugation
speed was essential to apply a uniform layer of cells on the electrode.
Photoactivation treatments consisted of 3 min of 10-Hz flashes. To
diminish photoinhibition phenomena, a red filter (similar to RG-610)
was interposed between the sample and the lamp. The filter was removed
for the subsequent experiments to obtain a saturating intensity. The
typical protocol was to keep the cells in the dark for about 15 min,
during which time the sample was prepared and centrifuged onto the
electrode. The electrode was polarized just before the experiment, and
15 flashes were given at a 3-Hz frequency. These cells were then left
on the electrode for an additional 3 min, with flashes given at a 10-Hz
frequency without recording the signal. The cells were then briefly
left in the dark for 10 s, and flashes were given again at a 3-Hz
frequency.
Analysis of O2 Yields
We tried to analyze the O2 yields after dark
adaptation from the same experiment as shown in Figure 4 for the 36-h
period from 144 to 180 h. However, O2
evolution in Cyanothece sp. was much more complex than in
Synechocystis sp. PCC 6803. Applying the correction
previously defined for the slow respiratory O2 transients (Meunier et al., 1996) produced worse fittings, since part
of the slow signals oscillated with a periodicity of 4 and was of PSII
origin (Fig. 1). Moreover, large corrections (up to 5 times greater
than those used with Synechocystis sp.) were necessary to
completely remove the slow O2 yields from the
data set in the Cyanothece sp. experiments. Therefore, we
used the averaged, smaller correction applied to
Synechocystis sp. PCC 6803 data (ratio of 3.0; see Meunier
et al., 1995a). This produced the best agreement between the
Cyanothece sp. data and S-state models.
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| Figure 4.
O2 yields from
Cyanothece sp. ATCC 51142 using cells harvested at L12
to D6 during growth under N2-fixing conditions in 12-h light/12-h dark conditions. The samples were stimulated by Xe flashes
given at 3 Hz starting 0.1 s after the start of recording. Since
the L12 to D0 boundary involved a change in culture flasks, the
synchronicity of the two cultures at that point was verified by
performing both recordings (12 h apart). The two recordings are
superimposed on the graph and are indistinguishable. An arbitrary constant was added to the recordings for readability (a.u., arbitrary unit).
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| Figure 1.
O2 yields from
Cyanothece sp. ATCC 51142 using cells harvested at L0 to
L4 during growth under N2-fixing conditions in 12-h light/12-h dark conditions. The samples were stimulated by Xe flashes
given at 3 Hz starting 0.1 s after the start of recording. An
arbitrary constant was added to the recordings for readability (a.u.,
arbitrary units).
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The O2-yield data after dark adaptation were
analyzed as described in the appendix of Meunier et al. (1996) to find
the five eigenvalues relating to a generic five-step S-state mechanism. The photoactivation probability, , was calculated from the first (greatest) eigenvalue,  1, as
1  1. This was a very reliable determination
for the following reasons: (a) 1 has the
lowest uncertainty (least sensitivity to experimental error) by a
factor of approximately 10 compared with the other eigenvalues; (b) its determination did not depend on any specific model; and (c) the only
assumption was that the data could be fitted by a five-step model.
Although the other eigenvalues were consistent with our cyanobacterial
model (Meunier et al., 1996), the low amplitudes of O2 evolution and the low signal-to-noise ratio
after dark adaptation made the determination of the other S-state
transition probabilities ambiguous at times, especially the separation
between "true" misses and the effect of deactivations. We found
from the simulation of theoretical sequences that the signal-to-noise
ratio needed to be significantly greater than 100 (1% error) for
fitting this model to data. However, the signal-to-noise ratio after
dark adaptation was often at or below 100. Therefore, we did not use
the cyanobacterial S-state model (Meunier et al., 1996) in this study.
Heterogeneous Photoactivation Model
The O2-yield data after a photoactivation
treatment (3 min of 10-Hz flashes with a red filter similar to RG-610)
had much greater amplitudes and provided a high signal-to-noise ratio
(on average around 400). Nevertheless, the data were much better fitted by a generic six-step model equation than with a four- or five-step one
(Meunier et al., 1995b). This was puzzling since there should not have
been a significant number of centers in S2 and S1 after the photoactivation treatment. The
30-s dark adaptation between the photoactivation treatment and the
following measurements were also unlikely to yield a significant number
of centers in S2 and S1
at all times, since even S3 was still present in
large amounts. We found that the requirement for a six-step analysis
was much better explained by the combination of two heterogeneous
four-step S-state models than by super-reduced S-states. Equation 8 in
Meunier et al. (1996) was modified for a six-step mechanism by adding
the term
c66n-1:
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(1)
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where the symbols have the same meanings as previously described.
Fitting Equation 1 resulted in a numerically more robust determination
of the eigenvalues than searching for roots in the characteristic
equation after multivariable regression (as described in the appendix
of Meunier et al., 1996). The eigenvalues 3, 4, 5, and
6 were all imaginary numbers. No consistent
six-step S-state model could reproduce these eigenvalues. However, the properties of the O2 yields could be
satisfactorily explained by the combination of two four-step S-state
models:
|
(2)
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where d and  serve the same function as c
and in the second four-step S-state model. Moreover, when
1 and 2 are close enough to 1 and 2,
Equation 2 provides an exact solution to interpreting Equation 1 as the
sum of two S-state models:
and where f1 + d1 = c1 and
f2 + d2 = c2. From these two sets of four
eigenvalues, the S-state transition probabilities of two four-step
homogeneous S-state cycles with backward transitions were calculated
(Meunier, 1993). This model is derived from a previously published
S-state model (Kok et al., 1970), in which the apparent misses
(failures to advance the S-state mechanism before the next flash) do
not distinguish between true photochemical misses or single hits
followed by deactivations. The possible occurrence of a miss followed
by a deactivation is accounted for by the backward transition
probability, so the backward transition probability is always smaller
than the miss probability.
The exact number of centers that contributed to each cycle is
proportional to the constants f1 and
d1, whereas only their sum
(c1) could be determined. However, most of
the time the variables c5 and
c6 were large enough relative to
c3 and c4 to
estimate the proportion of centers in the second (inefficient) cycle to more than 10%. Theoretical sequences generated by adding two four-step S-state cycles were tested as described previously (Meunier et al.,
1995b) and were found to also require a six-step model for analysis. By
contrast, the bicycle model (Shinkarev and Wraight, 1995) required only
a four-step model that returned the averaged properties of the
two theoretical cycles.
Fluorescence Measurements
PAM fluorescence was measured using the Walz fluorometer
ED-101 cuvette kit (Heinz-Walz, Effeltrich, Germany) with the optional 590-nm excitation light (model L-590-102, Heinz-Walz), and by adding a
small magnet and stirrer to keep the sample agitated. In the
experiments reported here, the actinic light was the original red LED
that came with the PAM instrument, and had an intensity of 68 µE
m2 s1. Transitions to
state 1 were induced by adding 10 µm DCMU and illuminating the samples with the red actinic light. The final FM(D1) level (D for DCMU and 1 for state 1)
was measured after the state transition was completed, as indicated by
a stabilization of the fluorescence level. It was verified previously
that this treatment induced substantial changes in 77 K fluorescence
(Meunier et al., 1997). However, because (a) state transitions in
cyanobacteria involve important changes in energy distribution; (b) the
resting state in the dark of Cyanothece sp. ATCC 51142 can
be anything between state 2 and state 1 (Meunier et al., 1997); and (c)
we used 590-nm excitation flashes for the detection of fluorescence, the FO level measured in
Cyanothece sp. ATCC 51142 was highly dependent on state
transitions. Therefore, we contend that the FM level obtained after a transition to
state 1 cannot be compared with the FO
level measured in dark-adapted "resting" cyanobacteria, FO(R), because phycobilisome attachment and
energy distribution changed. Therefore, after reaching state 1 the
fluorescence level (after a sufficient time for the reoxidation of
QA in darkness) FO(D1) is higher than
FO(R) because more energy is directed to PSII centers. Consequently, the fluorescence obtained per detecting flash of the PAM instrument is higher. It follows that
FM(D1) should be compared with
FO(D1) (Fig. 6).
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| Figure 6.
PAM fluorescence monitoring of the induction of
state 1 in Cyanothece sp. ATCC 51142 at D2 and D3. The
fluorescence levels used in calculations are tagged as described in
``Materials and Methods''. R, D, F, A, and 1, resting state in the
dark, in the presence of DCMU, measurements with a saturating flash,
measurements under actinic light, and state 1, respectively. Samples of
1 mL at 2 µg chlorophyll/mL were treated with 10 µm
DCMU and the maximum intensity of the red actinic light of the PAM
fluorimeter to induce state 1, as per the protocol in Meunier et al.
(1997). The actinic light went off just before the flash.
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We noticed that the difference FM(D1) FO(D1) after the treatment to induce a
transition to state 1 was smaller at some times during the cycle, even
though the samples were all at the same chlorophyll concentration. We
attributed the smaller difference to a residual
qN, calculated using the maximum
difference, MAX[FM(D1) FO(D1)], measured during the cycle as the
denominator:
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(D1)</UP></SUB>−F<SUB><UP>O(D1)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(D1)</UP></SUB>−F<SUB><UP>O(D1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img006.gif)
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(3)
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by contrast to the total qN,
calculated as:
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(DR)</UP></SUB>−F<SUB><UP>O(DR)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(D1)</UP></SUB>−F<SUB><UP>O(D1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img007.gif)
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(4)
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where FM(DR) is the
FM level measured in the presence of DCMU
(D) in the dark resting state (R) of the cyanobacterium,
FO(DR) is the
FO level measured in the presence of DCMU
in the dark-resting state of the cyanobacterium,
FO(D1) is the
FO level measured in the presence of DCMU
in state 1, and FM(D1) is the
FM level measured in the presence of DCMU
in state 1. The FO level was not measurably changed by the addition of DCMU, showing that the detecting light flashes had a negligible actinic effect; however, for the sake of
consistency, all levels were measured in the presence of DCMU. We found
two ways to measure the FM(D1) level: with
a saturating flash (F), FM(FD1), or with
the actinic light (A), FM(AD1). Likewise, the FM(DR) could be measured with a flash
(F), FM(FDR), or with the actinic light,
FM(ADR) (Fig. 6). This gives rise to four
ways to calculate qN:
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(FD1)</UP></SUB>−F<SUB><UP>O(FD1)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(FD1)</UP></SUB>−F<SUB><UP>O(FD1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img008.gif)
|
(5a)
|
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(AD1)</UP></SUB>−F<SUB><UP>O(AD1)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(FD1)</UP></SUB>−F<SUB><UP>O(FD1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img009.gif)
|
(5b)
|
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(FDR)</UP></SUB>−F<SUB><UP>O(FDR)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(D1)</UP></SUB>−F<SUB><UP>O(D1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img010.gif)
|
(5c)
|
![q<SUB><UP>N</UP></SUB>=1−(F<SUB><UP>M(ADR)</UP></SUB>−F<SUB><UP>O(ADR)</UP></SUB>)/<UP>MAX</UP>[F<SUB><UP>M(D1)</UP></SUB>−F<SUB><UP>O(D1)</UP></SUB>]](/content/vol116/issue4/fulltext/1551/img011.gif)
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(5d)
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RESULTS |
Metabolic Rhythms
The physiology of Cyanothece sp. ATCC 51142 has been
detailed previously for cells grown under
N2-fixing conditions with 12-h light/12-h dark or
with continuous light (Schneegurt, et al., 1994;
Colón-López, et al., 1997; Meunier, et al., 1997). In this
study, cells were grown in 12-h light/12-h dark cycles. In such
cultures nitrogenase and respiration activity peaked around D4, whereas
O2 evolution peaked near L8. Samples were
withdrawn every hour for the O2-flash yield or
the PAM measurements to be described. We will define four main time
periods: (a) early dark (D0 to the peak of nitrogenase activity); (b)
late dark (D6-D12); (c) early light (L0-L6); and (d) late light
(L8-L12). Although each procedure was repeated at least three times
with similar results, we will present results from single experiments
because the transitions are very sharp but the exact time point when
they happen varies from experiment to experiment (therefore, averaging blurs the transitions).
O2 Evolution
Cyanothece sp. ATCC 51142 produced
O2 under flashing light with a period-4
oscillation indicative of the S-state mechanism (Fig.
1). Samples harvested at various times
showed that the S-state mechanism underwent significant changes
throughout the 24-h cycle. Samples harvested at L0, the beginning of
light growth, showed slow O2 signals, peaking
about 45 ms after the flash, with oscillations period 4 in amplitude.
However, samples harvested at later times showed increasingly faster
O2 signals (peaking at 8 ms after the flash at
4 h of light) oscillating with a period 4, whereas the contribution of the slow transients to the oscillations became progressively less important.
In Synechocystis sp. PCC 6803, slow O2
signals were attributed to respiratory transients due to PSI activity
(Meunier et al., 1995a). These signals persisted in the presence of the
PSII herbicide DCMU, did not oscillate with a periodicity of 4, occurred in mutants with incapacitated PSII, and the deconvolution of
the fast O2 transients from the slow signals
resulted in greatly improved fittings with generic n-step models
(Meunier et al., 1995a). Attempts to completely remove the slow signals
from the data (as described in Meunier et al., 1995a) did not result in
improved fittings for Cyanothece sp. ATCC 51142 data. Slower
PSII O2 release has been reported in the
psbO deletion mutant of Synechocystis sp. PCC
6803 (Burnap et al., 1992). In light of the
qN data (see later), the slow
O2 release at the beginning of the light period
and increasingly faster O2 signals thereafter
would be consistent with a nonoptimal conformation of the oxidizing
side of PSII initially, and a subsequent photoactivation of PSII
centers.
The amplitudes of the O2 yields after sample
preparation in the dark increased from L0 to a maximum around L4 to L6;
however, they started decreasing past L6 down to a minimum between L8
and L10, and recovered quickly afterward (Fig.
2). The dip in
O2-evolution amplitudes in the middle of the
afternoon was unexpected, given the rates of linear electron transport
that have been observed (Meunier et al., 1997). However, a
photoactivation treatment of 3 min of 10-Hz flashes followed by 30 s of darkness produced very high amplitudes, with very strong period-4
oscillations; an example of this phenomenon for L10 is presented in
Figure 3. The effectiveness of the
photoactivation treatment demonstrated that the apparent loss of
activity from L8 to L10 after dark adaptation was reversible. Note that
the amplitudes obtained after dark adaptation in Cyanothece sp. ATCC 51142 (Fig. 2) were much lower than the actual PSII capacity (e.g. Fig. 3), and the amount of inactivation in the dark was variable
between repeats, whereas the trends remained the same. A striking
demonstration of the highly variable dark-inactivation properties of
Cyanothece sp. ATCC 51142 is shown in Figure 8.
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| Figure 2.
O2 yields from
Cyanothece sp. ATCC 51142 using cells harvested at L6 to
L12 during growth under N2-fixing conditions in 12-h light/12-h dark conditions. The samples were stimulated by Xe flashes
given at 3 Hz starting 0.1 s after the start of recording. An
arbitrary constant was added to the recordings for readability (a.u.,
arbitrary units).
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| Figure 3.
O2 yields from
Cyanothece sp. ATCC 51142 at L10 after a dark adaptation
(D) and a photoactivation treatment (P). The dark period necessary to
prepare the sample and centrifuge the cells to the electrode (about 15 min) was sufficient to deactivate most Mn centers (D). The same cells
were left on the electrode and photoactivated by 3 min of flashes given
at 10 Hz. After 10 s of darkness, flashes were given at 3 Hz and
the amplitude of the O2 yields were recorded (P). An
arbitrary constant was added to the recordings for readability (a.u.,
arbitrary unit).
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| Figure 8.
Amplitudes of O2 production under
flashing light from Cyanothece sp. ATCC 51142 cells
harvested at L8 (A) and D8 (B). A, Curve 1, after controlling light
leakages during centrifugation and sample manipulation ( ); curve 2, after 30 s of darkness following A ( ); curves 3, 4, and 5, each
after an additional 30 s of darkness following the previous
experiment ( ,  , and  ); curve P, after a 3-min, 10-Hz
photoactivation treatment following the experiment ( ). B, After
controlling light leakages during centrifugation and sample
manipulation (); after a subsequent 30 s of darkness ();
after an additional 30 s of darkness following the previous experiment ( and ).
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The amplitudes of the O2 yields after sample
preparation in the dark increased progressively from D0 (L12) to D2,
but decreased precipitously to negligible levels at D3 (Fig.
4; note the different scale from Fig. 2).
This corresponds to the usual time of the peak of nitrogenase activity
and respiration in Cyanothece sp. ATCC 51142 (Reddy et al.,
1993; Schneegurt et al., 1994; Colón-López et al., 1997).
Nitrogenase activity has a very narrow peak in 12-h light/12-h
dark-grown Cyanothece sp. ATCC 51142, and the mean for 27 cycles was at D4 (Colón-López et al., 1997). The amplitudes
of the O2 yields only partially recovered at D4
and later, suggesting that some PSII were inhibited during
N2 fixation. The transients measured from D7 to
D11 are very similar to those at D6 and, for clarity, are not shown.
To better understand these phenomena, flash-O2
yields were measured on cells harvested every hour for a 3-d period
using the photoactivation protocol used in Figure 3 (see Meunier et
al., 1997 for the metabolic periodicities). After a photoactivation treatment, the experimental data (Fig. 5,
solid line) showed a general trend that could be represented by a
24-h-period sine wave (dashed line). The photoactivated amplitudes on
the bare Pt electrode varied in a manner similar to the results
obtained using the Clark electrode (Meunier et al., 1997), and were
much higher than after dark adaptation. For this experiment, we
conclude that maximal activities are highest when cells are growing in the late-light phase (about L8) and lowest in the late-dark period.
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| Figure 5.
Average amplitudes of O2 production
under flashing light in Cyanothece sp. ATCC 51142 after
a photoactivation treatment and over a 3-d period starting 108 h
after subculture. Black bars, Dark periods; white bars, light periods.
The dashed line represents a sine wave with a fixed 24-h period, which
had the phase and amplitude fitted to the data using the DeltaGraph
program (Delta Point, Inc., Monterey, CA). a.u., Arbitrary units.
|
|
Fluorescence Quenching
The measurement of both 77 K and PAM fluorescence during
illumination of cells in the presence of DCMU indicated that several strong state transitions were occurring in Cyanothece sp.
ATCC 51142 throughout the 24-h cycles (Meunier et al., 1997).
qN was proposed to be mainly due to state
transitions (Campbell and Öquist, 1996). The calculation
qN uses the difference between the maximum level of fluorescence, FM, and the minimal
level, FO, under a test condition, compared
with the maximized difference in the presence of DCMU (see ``Materials and Methods''). However, there was an ambiguity in the evaluation of
the FM levels at certain times during the
cycle (Fig. 6; see ``Materials and Methods''). The fluorescence levels measured by using the red actinic
light of the PAM fluorimeter, FM(ADR) and
FM(AD1), were in practice equal to those
measured with a saturating flash, FM(FDR)
and FM(FD1), in wild-type
Synechocystis sp. PCC 6803 (not shown) and in
Cyanothece sp. ATCC 51142 at D2 (Fig. 6). However, these two
types of levels were significantly different from D3 (Fig. 6) until
approximately L4. This means that the maximum intensity of the actinic
red light in the PAM could not always maintain all of the
QA in a reduced state in Cyanothece
sp. ATCC 51142, even in the presence of DCMU.
We used the two ways to measure FM to
calculate the residual qN after inducing a
transition to state 1 (Fig. 7A).
According to Campbell and Öquist (1996), negligible
qN should remain in cyanobacteria after the
induction of state 1. However, the residual qN measured with the flash (Eq. 5a; Fig.
7A) increased in the dark (starting from D2), up to considerable levels
from D5 to D12, and decreased in the light. The amount of
qN that was not due to state transitions
was negligible from L5 to D1. By comparison, the residual
qN measured under red actinic light of the
PAM fluorimeter was greater from D4 to D12 than that measured by
flashes, which implies that it was harder to maintain
QA in its reduced form, and suggests a greater
instability of PSII charge separations (Eq. 5b; Fig. 7A). Indeed, in
PSII centers with an inactive donor side, excitation energy is quenched
by increased charge recombinations between QA
and P680+, which is observable
as an increased fluorescence quenching during the inactivation of
O2 evolution (Krieger et al., 1992). We interpret the qN not due to state transitions between
D4 and D12 as the result of an inhibition of PSII.
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| Figure 7.
Calculations of qN in 1 mL of Cyanothece sp. ATCC 51142 grown under 12-h
light/12-h dark conditions using the levels of fluorescence depicted in
Figure 6. Black bar, dark period; white bar, light period. A, Remaining
qN after the induction of state 1 measured with flashes ( ) or with the actinic light ( ). B, Total
qN in the dark in the presence of DCMU
measured with flashes () or with the actinic light ().
|
|
The total qN determined under flashes of
light (see ``Materials and Methods'') was important during the dark
period and decreased during the light period (Eq. 5c; Fig. 7B). The
fluorescence quenching determined from the red actinic light (Eq. 5d;
Fig. 7B) was identical, but only from L4 to D2; it was much higher from
D4 to D12. The high fluorescence quenching from D4 to D12 was
inconsistent with the previously reported state transitions, especially
the strong state 1 observed at D6 (Meunier et al., 1997). This, plus
the lower amplitudes of O2 evolution under
flashing light, suggest that PSII centers were inactivated or inhibited
in the dark during N2 fixation. The disappearance
of that source of quenching in the light is consistent with a
subsequent photoactivation or repair of PSII centers. It follows that
the inactivation and photoactivation mechanisms in
Cyanothece sp. ATCC 51142 are of particular interest.
The inactivation and photoactivation processes in Cyanothece
sp. ATCC 51142 demonstrated unusual properties and striking differences between cells harvested at L8 and D8. Cells at L8 and D8 were harvested
and kept in absolute darkness from the time of harvest until treatment;
these precautions preserved activity. L8 cells illuminated with 3-Hz
flashes produced an O2-evolution pattern shown in
Figure 8A. A sample was then left on
the electrode in the dark for 30 s, after which 15 additional
flashes were given. Under these conditions, PSII
O2 evolution declined more than 2-fold (Fig. 8A,
curve 2). This pattern could be repeated several times until very
little PSII activity remained (Fig. 8A, curves 3-5). This loss was
reversible, as demonstrated by a 3-min photoactivation treatment of
10-Hz flashes prior to a flash train (Fig. 8A, curve P). The sample was
in the dark for 15 min before these experiments and thus remained
stable in complete darkness. This demonstrated that the dark
inactivation of PSII was stimulated by exposure to 3-Hz light flashes
followed by dark incubation. This property, coupled with the metabolic
rhythms, made the determination of the decay times of
S3 and S2 impractical (at
least near L6-L10).
By contrast, cells harvested at D8 were relatively insensitive to the
same treatment (Fig. 8B). After the initial measurement, PSII activity
remained approximately constant after many cycles of light flashes and
dark incubation. This experiment demonstrated that the effectiveness of
3-Hz flashes in triggering dark losses of activity was negligible in
cells harvested at D8. However, it must be noted that the amplitudes of
the O2 yields were about 3-fold lower in D8 cells
than in L8 cells. This is one of the compelling reasons to conclude
that many PSII centers were inhibited in the dark-grown cells.
Analysis of O2 Yields
As discussed in "Materials and Methods," the low
O2 yields observed after dark adaptation resulted
in a poor signal-to-noise ratio that prevented the unambiguous and
reproducible fitting of S-state models. However, the photoactivation
probability could be reliably recovered since it was calculated
directly from the robust first eigenvalue, independently of any other
probabilities or model assumptions (see ``Materials and Methods'').
The photoactivation of centers, , showed three repeating peaks, P1
to P3 (Fig. 9). P1 was close to the
peak of N2 fixation, whereas P2 and P3 were in
the first half and the second half of the light phase, respectively. Of these photoactivation peaks, only P2 was correlated with an increase in
PSII activity. Clearly, the photoactivation probability after dark
adaptation reflected a phenomenon other than an increase in the
steady-state level of active PSII centers. However, the photoactivation
treatment described in ``Materials and Methods'' was effective in
lowering to 0, suggesting that could be inversely correlated
with the dark stability of PSII. Indeed, P3 correlated with the
reversible high dark instability of PSII (Figs. 2, 3, and 8), and P1
correlated with the inactivation of PSII centers.
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| Figure 9.
Photoactivation probability, , in dark-adapted
cells of Cyanothece sp. ATCC 51142 for the last 48 h of the experiment shown in Figure 5 (). Black bars, dark periods;
white bars, light periods. P1, P2, and P3, Significant repeating
periods of high photoactivation; the location of the peaks should be
taken as the center of mass of the area under the curve (which is
effectively spaced 24 h apart, whereas the maximum measured
photoactivation probability is not necessarily spaced this way). The
modeling errors were represented by the standard deviations calculated
from the error matrix returned by the multivariable linear regression
over a generic five-step equation. They were found to be smaller than or equal to 0.1% + 0.1 × , which are the values shown by the error bars. As many O2 yields as possible were used for the
regression (in most cases, 14); however, the results were insensitive
to the number of flashes used and to the inclusion of the first flash. The results were essentially identical to a generic four-step equation
ignoring the first flash.
|
|
The O2 yields after photoactivation in
Cyanothece sp. ATCC 51142 proved impossible to analyze
consistently in the framework of variants of single Kok models (not
shown). The six-step eigenvalue analysis of photoactivated
O2 yields in Cyanothece sp. ATCC 51142 (Meunier et al., 1995b) provided a clue as to the origin of the problem
by always finding two pairs of conjugated eigenvalues instead of the
usual single pair. The experimental eigenvalues found for samples
harvested at all times had a first eigenvalue equal to 1; therefore,
the sum of all probabilities was 1 and the photoactivation probability
was 0. This indicated that the photoactivation treatment had been
effective in removing quickly photoactivatable PSII centers. The
second, real and negative eigenvalue related to the damping mechanisms
(Meunier, 1993) was also present within the usual range of values.
However, two conjugated pairs of eigenvalues with large imaginary
components were found instead of one conjugated pair and two more real
eigenvalues. By mathematical necessity, the eigenvalues obtained from a
Kok model (even extended to S2) always contain
only one pair of conjugated eigenvalues because there is only one
cycle.
The two conjugated pairs suggested the presence of two S-state cycles
(in different PSII centers) with significantly different properties.
The consideration of PSII heterogeneity in the O2 yields permitted the successful interpretation of the eigenvalues (Table I). The combination of a
dissipative or inefficient cycle with an efficient one could reproduce
the O2 yields down to the level of experimental
noise. The efficient centers retained similar properties, with about 9 to 11% apparent misses, 2 to 4% double-hits, and 0 to 1% backward
transitions. By contrast, the inefficient centers changed throughout
the cycle in number and in properties. The inefficient centers were
characterized by a lower double-hit probability and higher backward
transition and miss probabilities. The inefficiency was the most benign
around D5, close to the previously observed strong state 1 and
(presumed) oxidized state of the plastoquinone pool (Meunier et al.,
1997).
View this table:
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|
Table I.
Calculated S-state transition probabilities for the
two heterogeneous S-state cycles at different times during the
metabolic rhythms of Cyanothece sp. ATCC 51142
The single-hit probability is equal to 1 minus the sum of the other
probabilities because after the photoactivation treatment, the first
eigenvalue was always very close to 1.
|
|
There were too few inefficient centers to quantify their properties at
L5, in agreement with our proposal that centers were photoactivated in
the early light period. The numbers provided in Table I are not unique,
because there are many combinations of heterogeneous cycles that could
produce O2 yields close to the experimental ones.
However, this interpretation is the simplest, being an exact solution,
and is consistent throughout the metabolic rhythms in
Cyanothece sp. ATCC 51142. Thus, PSII heterogeneity could
explain the properties of the O2 yields found
after photoactivation. We interpret this as evidence for a difference
in the properties of freshly activated PSII (inefficient PSII) compared
with the centers that were stable in the dark.
| |
DISCUSSION |
The data presented in this paper indicated that
O2 evolution underwent many changes during the
diurnal cycle of the diazotrophic cyanobacterium Cyanothece
sp. ATCC 51142. The capacity for O2 evolution,
the time required for O2 release by the S-state
mechanism, and the dark stability of PSII centers changed dramatically
throughout a diurnal cycle. In particular, the data reflect a
substantial level of PSII heterogeneity as the cells are preparing for
N2 fixation. It is evident that PSII is a main
target for down-regulation during nitrogenase activity.
The regulation of photosynthesis in Cyanothece sp. ATCC
51142 is mediated in part through state transitions and changes in reaction-center oligomerization (Meunier et al., 1997). The source of
qN that developed in the early dark was an
inhibition of PSII that lowered variable fluorescence
(Fv [= FM FO]), even in the presence of DCMU, and
which led to lower amplitudes of O2 evolution. The dark inactivation of PSII centers during N2
fixation occurred while PSII centers were in the monomeric form
following a transition to state 2 (Meunier et al., 1997). After dark
adaptation, the amplitudes of O2 evolution were
negligible at D3; however, the amplitudes after 3 min of 10-Hz flashes
were comparable to one-third of the maximum PSII amplitudes. The
partial recovery may be explained by a reduced state of the
plastoquinone pool during the intense respiratory activity associated
with N2 fixation and a re-oxidation of the
plastoquinone pool by the flashes. The fact that it was partial
demonstrates that some PSII were inhibited. Indeed, the previously
observed sudden and strong transition to state 1 after the peak in
N2 fixation, indicating an oxidation of the
plastoquinone pool, appeared correlated with only a partial recovery of
O2 evolution in the experiments reported here.
Meanwhile, the PAM fluorescence method was unable to detect state
transitions in the period from D3 to L1, due to the significant
inhibition of PSII.
The photoactivation of PSII centers in the early light occurred while
the cells were close to state 2, which favors excitation of PSI.
Progressively, as PSII centers were photoactivated, the qN diminished and revealed a strong state 1 toward the end of the light period. However, at any given time, the PAM
method was ambiguous as to the contributions to
qN from state transitions or from
not-yet-photoactivated centers. The analysis of the
O2 yields revealed a strong photoactivation
probability in the early part of the light phase, whereas the 77 K
spectra in Meunier et al. (1997) indicated that most of the transition
to state 1 occurred in the second half of the light phase. Thus, the
photoactivation of PSII centers was prevalent in the first half of the
light phase, when the centers were mainly in monomeric form.
At the beginning of the light period, when most PSII centers were
inhibited, the S-state mechanism produced slow O2
signals of a period 4. As photoactivation progressed, these changed to fast signals (Figs. 1 and 2), demonstrating the simultaneous presence of PSII centers with differing properties that were interconvertible. The O2 produced was contributed by (at least) two
populations of PSII centers and required two S-state cycles for
modeling. The data we obtained for the interconversion and
photoactivation processes suggests the repair of an inefficient,
dark-unstable PSII form to an efficient, faster one.
We have developed a modified model of the PSII S-state mechanism in
cyanobacteria (Meunier et al., 1996), which has formalized the
significant differences in flash-yield experiments between cyanobacteria and chloroplasts. The modeling of cyanobacterial O2 evolution required explicit deactivations in
the dark interval between flashes and in homogeneous probabilities, and
an explicit accounting of the number of active PSII centers by
photoinhibition or photoactivation (Meunier et al., 1996). The model
was used to explain some of the interesting properties of
Synechocystis sp. PCC 6803 wild type and the psbO
deletion mutant (lacking the Mn2+-stabilizing
protein MSP) (Meunier et al., 1996).
The comparison of flash-yield results among Synechocystis
wild type, the psbO deletion mutant, and
Cyanothece cells from different times in the diurnal cycle
led to the discovery of some important similarities.
O2 evolution by wild-type
Synechocystis sp. PCC 6803 is very stable in the dark and
remains so even after a prolonged period of centrifugation onto a bare
Pt electrode (Burnap et al., 1996; Meunier et al., 1996). The
psbO deletion mutant of Synechocystis sp. PCC
6803 shows very poor O2 evolution after dark
incubation. However, after a 3-min flash regime at 10 Hz, the same
sample showed a greatly enhanced production of O2
(i.e. high photoreactivation; Meunier et al., 1996, fig. 3). Therefore,
we suggest that Cyanothece sp. ATCC 51142 cells sometimes
during the light period have an O2-evolving
mechanism that resembles the psbO deletion mutant of
Synechocystis sp. PCC 6803. At the same time, the PSII
centers of Cyanothece sp. cells growing in the dark are
fewer, but possess a dark stability more similar to that of
Synechocystis sp. PCC 6803 wild type. Moreover, 77 K spectra
indicated changes in the oligomerization of PSII centers (Meunier et
al., 1997). These observations and the replacement of the D1 protein
suggest that the protein conformations, composition, and
oligomerization of PSII centers are under constant flux in
Cyanothece sp. ATCC 51142.
| |
FOOTNOTES |
1
This work was supported by grants from the U.S.
Department of Energy (no. DE-FG02-89ER14028A) and the U.S. Department
of Agriculture (no. 93-37306-9238) to L.A.S.
*
Corresponding author; e-mail
lsherman{at}bilbo.bio.purdue.edu; fax
1-765-496-1495.
Received September 22, 1997;
accepted December 18, 1997.
| |
ABBREVIATIONS |
Abbreviations:
DX, X h of darkness.
FM, maximum fluorescence.
FO, initial fluorescence.
LX, X h of light.
PAM, pulse-amplitude-modulated.
qN, nonphotochemical quenching.
| |
ACKNOWLEDGMENT |
We are grateful to Dr. Rob Burnap of Oklahoma State University
for the use of his bare Pt electrode equipment and for his suggestion
to photoactivate the cells.
| |
LITERATURE CITED |
Arnon DI,
McSwain BD,
Tsujimoto HY,
Wada K
(1974)
Photochemical activity and components of membrane preparations from blue-green algae I: coexistence of two photosystems in relation to chlorophyll a and removal of phycocyanin.
Biochim Biophys Acta
357:
231-245
[Medline]
Bader KP
(1994)
Physiological and evolutionary aspects of the H2O/H2O2-cycle in cyanobacteria.
Biochim Biophys Acta
1188:
213-219
[CrossRef]
Bader KP,
Thibault P,
Schmid GH
(1983)
A study on oxygen evolution and on the S-state distribution in thylakoid preparations of the filamentous blue-green alga Oscillatoria chalybea.
Z Naturforsch
38c:
778-792
Bergman B,
Gallon JR,
Rai AN,
Stal LJ
(1997)
N2 fixation by non-heterocystous cyanobacteria [review].
FEMS Microbiol Rev
19:
139-185
[CrossRef][Web of Science]
Burnap RL,
Qian M,
Pierce C
(1996)
The manganese-stabilizing protein of photosystem II modifies the in vivo deactivation and photoactivation kinetics of the H2O oxidation complex in Synechocystis sp. PCC 6803.
Biochemistry
35:
874-882
[CrossRef][Medline]
Burnap RL,
Shen JR,
Jursinic PA,
Inoue Y,
Sherman LA
(1992)
Oxygen yield and thermoluminescence characteristics of a cyanobacterium lacking the manganese-stabilizing protein of photosystem II.
Biochemistry
31:
7404-7410
[CrossRef][Medline]
Campbell D,
Öquist G
(1996)
Predicting light acclimation in cyanobacteria from nonphotochemical quenching of photosystem II fluorescence, which reflects state transitions in these organisms.
Plant Physiol
111:
1293-1298
[Abstract]
Cheniae GM,
Martin IF
(1972)
Effects of hydroxylamine on photosystem II. II. Photoreversal of the NH2OH destruction of O2 evolution.
Plant Physiol
50:
87-94
[Abstract/Free Full Text]
Colón-López MS,
Sherman DM,
Sherman LA
(1997)
Transcriptional and translational regulation of nitrogenase in light-dark- and continuous-light-grown cultures of the unicellular cyanobacterium, Cyanothece sp. ATCC 51142.
J Bacteriol
179:
4319-4327
[Abstract/Free Full Text]
Engels DH,
Lott A,
Schmid GH,
Pistorious EK
(1994)
Inactivation of the water-oxidizing enzyme in manganese stabilizing protein-free mutant cells of the cyanobacteria Synechococcus PCC 7942 and Synechocystis PCC 6803 during the dark incubation and conditions leading to photoactivation.
Photosynth Res
42:
227-244
[CrossRef]
Fay P
(1992)
Oxygen relations of nitrogen fixation in cyanobacteria.
Microbiol Rev
56:
340-373
[Abstract/Free Full Text]
Fay P,
Cox RM
(1967)
O2 inhibition of N2 fixation in cell-free preparations of blue-green algae.
Biochim Biophys Acta
143:
562-569
[Medline]
Gleiter HM,
Haag E,
Shen JR,
Eaton-Rye JJ,
Seeliger AG,
Inoue Y,
Vermaas WFJ,
Renger G
(1995)
Functional characterization of mutant strains of the cyanobacterium Synechocystis sp. PCC 6803 lacking short domains within the large, lumen-exposed loop of the chlorophyll protein CP-47 in photosystem II.
Biochemistry
34:
6847-6856
[CrossRef][Medline]
Grobbelaar N,
Lin HY,
Huang TC
(1987)
Induction of a nitrogenase activity rhythm in Synechococcus and the protection of its nitrogenase against photosynthetic oxygen.
Curr Microbiol
15:
29-33
Haystead A,
Robinson R,
Stewart WDP
(1970)
Nitrogenase activity in extracts of heterocystous blue-green algae.
Arch Mikrobiol
75:
235-243
Kok B,
Forbush B,
McGloin M
(1970)
Cooperation of charges in photosynthetic O2 evolution-1. A linear four step mechanism.
Photochem Photobiol
11:
457-475
[Web of Science][Medline]
Kretschmann H,
Witt HT
(1993)
Chemical reduction of the water splitting enzyme system of photosynthesis and its light-induced reoxidation characterized by optical and mass spectrometric measurements: a basis for the estimation of the states of the redox active manganese and of water in the quaternary oxygen-evolving S-state cycle.
Biochim Biophys Acta
1144:
331-345
[CrossRef]
Krieger A,
Moya I,
Weis E
(1992)
Energy-dependent quenching of chlorophyll-A fluorescence: effect of pH on stationary fluorescence and picosecond-relaxation kinetics in thylakoid membranes and photosystem II preparations.
Biochim Biophys Acta
1102:
167-176
[CrossRef]
Messinger J,
Renger G
(1993)
Generation, oxidation by the oxidized form of the tyrosine of polypeptide D2, and possible electronic configuration of the redox states S0, S1, and S2 of the water oxidase in isolated spinach thylakoids.
Biochemistry
32:
9379-9386
[CrossRef][Medline]
Messinger J,
Seaton G,
Wydzynski T,
Wacker U,
Renger G
(1997)
S3 state of the water oxidase in photosystem II.
Biochemistry
36:
6862-6873
[CrossRef][Medline]
Meunier PC
(1993)
O2 evolution by photosystem II: the contribution of backward transitions to the anomalous behaviour of double-hits revealed by a new analysis method.
Photosynth Res
36:
111-118
[CrossRef]
Meunier PC,
Burnap RL,
Sherman LA
(1995a)
Interaction of the photosynthetic and respiratory electron transport chains producing slow O2 signals under flashing light in Synechocystis sp. PCC 6803.
Photosynth Res
45:
31-40
[CrossRef]
Meunier PC,
Burnap RL,
Sherman LA
(1996)
Improved 5-step modeling of the photosystem II S-state mechanism in cyanobacteria.
Photosynth Res
47:
61-76
[CrossRef]
Meunier PC,
Colón-López MS,
Sherman LA
(1997)
Temporal changes in state transitions and photosystem organization in the unicellular, diazotrophic cyanobacterium Cyanothece sp. ATCC 51142.
Plant Physiol
115:
991-1000
[Abstract]
Meunier PC,
Popovic R
(1988)
High-accuracy O2 polarograph for photosynthetic systems.
Rev Sci Inst
59:
486-491
[CrossRef]
Meunier PC,
Watters JW,
Colón-López MS,
Sherman LA
(1995b)
Regulation of the O2-evolving mechanism during N2 fixation in the diazotrophic cyanobacterium Cyanothece sp. ATCC 51142.
In
P Mathis,
eds, Research in Photosynthesis, Vol 2.
Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 389-392
Misra HS,
Desai TS
(1993)
Involvement of acceptor side components of PSII in the regulatory mechanism of Plectonema boryanum grown photoautotrophically under diazotrophic condition.
Biochem Biophys Res Com
194:
1001-1007
[CrossRef][Medline]
Misra HS,
Tuli R
(1994)
Nitrogen fixation by Plectonema boryanum has a photosystem II independent component.
Microbiology
140:
971-976
[Abstract/Free Full Text]
Mitsui A,
Cao S,
Takahashi A,
Arai T
(1987)
Growth synchrony and cellular parameters of the unicellular nitrogen-fixing marine cyanobacterium, Synechococcus sp. strain Miami BG043511 under continuous illumination.
Physiol Plant
69:
1-8
Miyao M,
Ikeuchi M,
Yamamoto N,
Ono T
(1995)
Specific degradation of the D1 protein of photosystem-II by treatment with hydrogen-peroxide in darkness: implications for the mechanism of degradation of the D1 protein under illumination.
Biochemistry
34:
10019-10026
[CrossRef][Medline]
Mullineaux PM,
Gallon JR,
Chaplin AE
(1981)
Acetylene reduction (nitrogen fixation) by cyanobacteria grown under alternating light dark cycles.
FEMS Microbiol
10:
245-247
[CrossRef]
Reddy KJ,
Haskell B,
Sherman DM,
Sherman LA
(1993)
Unicellular, aerobic N2-fixing cyanobacteria of the genus Cyanothece.
J Bacteriol
175:
1284-1292
[Abstract/Free Full Text]
Schneegurt M,
Sherman DM,
Nayar S,
Sherman LA
(1994)
Oscillating behaviour of carbohydrate granule formation and dinitrogen fixation in the cyanobacterium Cyanothece sp. strain ATCC 51142.
J Bacteriol
176:
1586-1597
[Abstract/Free Full Text]
Shinkarev VP,
Wraight CA
(1995)
Oxygen evolution in photosynthesis: from unicycle to bicycle.
Proc Natl Acad Sci USA
90:
1834-1838
[Abstract/Free Full Text]
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Top
Abstract
Introduction
