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A Soluble Carotenoid Protein Involved inPhycobilisome-Related Energy Dissipationin Cyanobacteria
Adjele Wilson,a Ghada Ajlani,b Jean-Marc Verbavatz,b Imre Vass,c Cheryl A. Kerfeld,d and Diana Kirilovskya,1
a Unite de Recherche Associee 2096, Centre National de la Recherche Scientifique, Service de Bioenergetique, 91191
Gif sur Yvette, Franceb Service de Biologie de Fonctionnes Membranaires, Departement de Biologie Joliot-Curie, Commissariat a
l’Energie Atomique Saclay, 91191 Gif sur Yvette, Francec Biological Research Center, H-6726 Szeged, Hungaryd Molecular Biology Institute, University of California, Los Angeles, California 90095-1570
Photosynthetic organisms have developed multiple protective mechanisms to survive under high-light conditions. In plants, one
of these mechanisms is the thermal dissipation of excitation energy in the membrane-bound chlorophyll antenna of photosys-
tem II. The question of whether or not cyanobacteria, the progenitor of the chloroplast, have an equivalent photoprotective
mechanism has long been unanswered. Recently, however, evidence was presented for the possible existence of a mechanism
dissipating excess absorbed energy in the phycobilisome, the extramembrane antenna of cyanobacteria. Here, we demonstrate
that this photoprotective mechanism, characterized by blue light–induced fluorescence quenching, is indeed phycobilisome-
related and that a soluble carotenoid binding protein, ORANGE CAROTENOID PROTEIN (OCP), encoded by the slr1963 gene in
Synechocystis PCC 6803, plays an essential role in this process. Blue light is unable to quench fluorescence in the absence of
phycobilisomes or OCP. The fluorescence quenching is notDpH-dependent, and it can be induced in the absence of the reaction
center II or the chlorophyll antenna, CP43 and CP47. Our data suggest that OCP, which strongly interacts with the thylakoids,
acts as both the photoreceptor and the mediator of the reduction of the amount of energy transferred from the phycobilisomes to
the photosystems. These are novel roles for a soluble carotenoid protein.
INTRODUCTION
Carotenoids, the most widely occurring pigments in nature, are
essential to microbial, animal, and plant life. These lipophilic
macromolecules play diverse roles, functioning as colorants,
precursors of visual pigments, and antioxidants. In plant and
algal photosynthesis, carotenoids are known to have a dual
function in chlorophyll–membrane complexes: harvesting light
and photoprotection. The photoprotective mechanisms include
(1) reducing the amount of energy funneled to photochemical
reaction centers (by screening or thermal energy dissipation), (2)
quenching triplet chlorophyll to prevent the formation of singlet
oxygen, and (3) directly quenching singlet oxygen.
In cyanobacteria, carotenoids are also associated with pro-
teins devoid of chlorophyll. These water-soluble carotenoid bind-
ing proteins were first described by Holt and Krogmann (1981)
and later found in several different cyanobacterial species (re-
viewed in Kerfeld, 2004a, 2004b). The soluble ORANGE CA-
ROTENOID PROTEIN (OCP), a 35-kD protein that contains a
single noncovalently bound carotenoid (Holt and Krogmann,
1981; Wu and Krogmann, 1997; Kerfeld, 2004a, 2004b), is the
best characterized of these proteins. In Synechocystis PCC
6803, the OCP is the product of the slr1963 open reading frame
(Wu and Krogmann, 1997). Highly conserved homologs of the
OCP are found in the genomes of all cyanobacteria for which
genomic data are available, with the exception of the prochlor-
ococci (Kerfeld, 2004a, 2004b).
The structure of the Arthrospira maxima OCP has been de-
termined to 2.1 A resolution (Kerfeld et al., 2003). The OCP
consists of two domains: an alla-helical N-terminal domain and a
mixed a/b C-terminal domain. The embedded carotenoid, a
39-hydroxyequinenone, has an all-trans configuration and spans
both protein domains. The protein has a large effect on the
spectroscopic characteristics of the carotenoid. In organic sol-
vents, the 39-hydroxyequinenone is yellow (lmax ¼ 450 nm), but
in the OCP, it appears orange (lmax ¼ 465 and 495 nm) (Kerfeld
et al., 2003; Polivka et al., 2005). In the course of OCP purifica-
tion, a red carotenoid protein (RCP) is also isolated (Wu and
Krogmann, 1997; Kerfeld, 2004a, 2004b). N-terminal sequencing
and mass spectrometry analysis indicated that this is a 16-kD
proteolytic fragment of the OCP. The proteolysis removes the
entire C-terminal domain, which, without concomitant structural
change, would expose nearly half of the carotenoid to solvent.
A role for the OCP under stress conditions has been proposed.
Indeed, microarray studies indicate that the OCP transcript
levels increase by >600% upon transfer to high light (Hihara
et al., 2001). Several functions for the OCP in photoprotection
have been suggested based on its structure and on in vitro
1 To whom correspondence should be addressed. E-mail [email protected]; fax 33-1-69088717.The authors responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy describedin the Instructions for Authors (www.plantcell.org) are: Ghada Ajlani([email protected]) and Diana Kirilovsky ([email protected]).Article, publication date, and citation information can be found atwww.plantcell.org/cgi/doi/10.1105/tpc.105.040121.
The Plant Cell, Vol. 18, 992–1007, April 2006, www.plantcell.org ª 2006 American Society of Plant Biologists
experiments, such as a quencher of singlet oxygen, a carotenoid
transport protein, and a light screen (Kerfeld, 2004a, 2004b).
By harvesting solar energy and converting it into chemical
energy, plants, algae, and cyanobacteria provide food and oxygen
that are essential for life on earth. However, excess light can be
lethal for photosynthetic organisms, because harmful reactive
oxygen species are generated in the photochemical reaction
centers when energy absorption exceeds the rate of carbon
fixation. To survive, photosynthetic organisms have evolved sev-
eral protective processes. In plants and algae, dissipation of the
excess absorbed energy as heat in the light-harvesting chlorophyll
antenna (LHCII) of photosystem II (PSII) decreases the amount of
energy funneled to the photochemical centers (for reviews, see
Demmig-Adams, 1990; Horton et al., 1996; Niyogi, 1999; Muller
et al., 2001). A decrease of PSII-related fluorescence emission,
known as nonphotochemical quenching (NPQ; more specifically
qE), is observed when this process is triggered by acidification of
the thylakoid lumen under saturating light conditions. A decrease
in thylakoid lumen pH activates the formation of the carotenoid
zeaxanthin from violaxanthin via the xanthophyll cycle (Yamamoto,
1979; Gilmore and Yamamoto, 1993). Low pH drives the proton-
ation of PsbS, a PSII subunit that belongs to the LHC superfamily
(Li et al., 2000), inducing conformational changes in the LHCII
(Ruban et al., 1992).
Cyanobacteria do not have the integral membrane chlorophyll-
containing light-harvesting complex, LHCII. Instead, light is cap-
tured by a membrane extrinsic complex, the phycobilisome,
which is attached to the outer surface of thylakoid membranes
(Gantt and Conti, 1966). These large complexes consist of
phycobiliproteins that covalently bind bilin pigments and linker
peptides that are required for the organization of the phycobili-
some and for tuning the physical characteristics of the pigments
(for reviews, see Glazer, 1984; MacColl, 1998; Tandeau de
Marsac, 2003; Adir, 2005). Phycobilisomes are composed of a
core from which rods (usually six) radiate. The major core protein
is allophycocyanin, whereas the rods contain phycocyanin and in
some species phycoerythrin or phycoerythrocyanin (in the distal
end of the rod). The phycobilisomes are bound to the thylakoids
via the core–membrane linker protein, Lcm, which also serves as
the terminal energy acceptor of the phycobilisomes (Redlinger
and Gantt, 1982). Harvested light energy is transferred from Lcm
to the chlorophylls of the inner chlorophyll antenna, CP43 and
CP47 (containing chlorophyll a and carotenoids), and to reaction
center II. Phycobilisome can also transfer energy to photosystem
I (PSI) (Mullineaux, 1992; Rakhimberdieva et al., 2001).
Cyanobacteria have two well-characterized mechanisms that
are associated with fluorescence quenching: state transitions
and photoinhibition. In photoinhibition, high light intensities in-
duce PSII fluorescence quenching and the irreversible inactiva-
tion of PSII caused by damage and degradation of the D1 protein,
an essential constituent of PSII (for reviews, see Prasil et al.,
1992; Aro et al., 1993; Melis, 1999). Recovery of fluorescence
and oxygen-evolving activity requires the replacement of the
damaged D1 protein. State transitions regulate the redistribution
of energy between PSII and PSI; they are induced by changes in
the redox state of the plastoquinone pool (for reviews, see Allen,
1992; van Thor et al., 1998; Wollman, 2001). Exposure of pho-
tosynthetic organisms to light absorbed predominantly by PSII
causes a relative decrease of the PSII fluorescence yield. Con-
versely, illumination with light absorbed preferentially by PSI
induces a relative increase of the fluorescence yield.
It has long been assumed that cyanobacteria do not use an
antenna-related NPQ mechanism to decrease the amount of en-
ergy funneled to reaction center II (Campbell et al., 1998). Several
recent experiments refute this view. For example, an NPQ mech-
anism mediated by the Iron stress–induced A protein (IsiA) that
belongs to the core complex family of chlorophyll binding pro-
teins has been described. This protein, which is induced under
iron starvation (Laudenbach and Straus, 1988; Burnap et al.,
1993) and other stress conditions (Jeanjean et al., 2003; Yousef
et al., 2003; Havaux et al., 2005), encircles the PSI reaction center
(Bibby et al., 2001; Boekema et al., 2001). After prolonged iron
starvation, empty rings of IsiA (without PSI) are also detected
(Yeremenko et al., 2004). It was proposed that this unconnected
IsiA is involved in photoprotection (Park et al., 1999; Sandstrom
et al., 2001; Yeremenko et al., 2004; Bailey et al., 2005; Ihalainen
et al., 2005). Indeed, it was demonstrated that these IsiA aggre-
gates are in a strongly quenched state, suggesting that they are
responsible for the thermal dissipation of absorbed energy
(Ihalainen et al., 2005).
In addition, results revealing the existence of a distinct blue
light–induced NPQ mechanism, proposed to be associated with
the phycobilisome, were first described in 2000 (El Bissati et al.,
2000). Subsequently, spectral and kinetic data were presented
suggesting that blue light–activated carotenoids induce the quench-
ing of phycobilisome fluorescence emission (Rakhimberdieva et al.,
2004).
Here, we report that the carotenoid protein OCP is essential for
this phycobilisome-associated NPQ mechanism in Synechocys-
tisPCC 6803. The OCP appears to act as both the photoreceptor
and the mediator of a photoprotective energy dissipation mech-
anism, which decreases the amount of energy arriving at both
photosystems from the phycobilisome.
RESULTS
Construction of OCP Mutants
To determine the function and cellular location of the OCP, we
constructed three mutants ofSynechocystisPCC 6803: a mutant
lacking the OCP protein, in which the ocp gene (slr1963) was
inactivated (DOCP); a mutant containing an OCP–green fluores-
cent protein fusion under the control of the ocp promoter (OCP-
GFP); and a mutant in which the gene slr1964 was inactivated
(DSlr1964), used as the control strain. An antibiotic resistance
cassette was introduced into the HincII site in slr1963 (DOCP) or
into the ClaI site in slr1964 (DSlr1964 and OCP-GFP) by double
homologous recombination (Figure 1A). To confirm the insertion
sites and the complete segregation of the mutants, PCR analysis
was performed. The amplification of the genomic region con-
taining slr1963, isolated from DOCP, DSlr1964, and OCP-GFP,
with the oligonucleotides car1 and car6 gave fragments of 4.0 kb
(DOCP and DSlr1964) and 4.8 kb (OCP-GFP) (Figure 1B). No
traces of the 2-kb fragment, observed in the wild type, were
detected in the mutants, indicating complete segregation. The
Carotenoids and NPQ in Cyanobacteria 993
correct position of the antibiotic cassette in DOCP and DSlr1964
was controlled by PCR amplification of slr1963 by the oligonu-
cleotides car1 and car2 (Figure 1B). In the DOCP mutant, the
3-kb PCR fragment indicated that slr1963 was interrupted by the
antibiotic cassette, whereas this gene was not interrupted in
DSlr1964 (0.9-kb PCR fragment). To check the position of the
antibiotic cassette in this mutant, the 4-kb PCR fragment ob-
tained by amplification with the oligonucleotides car1 and car6
was digested by the restriction enzyme HincII. As expected,
digestion of the DSlr1964 PCR fragment gave four fragments of
2 kb, 800 bp, 750 bp, and 440 bp, confirming that in this mutant
slr1964 is interrupted by the antibiotic cassette (Figure 1C).
Phenotype Conferred by DOCP under
High Light Intensities
We compared the behavior of DOCP and wild-type cells under
high-intensity white light. When fluorescence was measured with
a PAM fluorometer during saturating light illumination, a faster
quenching of fluorescence was observed in wild-type cells than
in DOCP cells. Figure 2 shows room temperature fluorescence
traces in Synechocystis wild-type and DOCP cells illuminated by
different light intensities. Dark-adapted wild-type and DOCP
cells presented a characteristic low level of maximal fluores-
cence (Fm). Upon illumination with low intensities of white light, a
maximal level of Fm was reached in both strains (Figure 2A). This
increase of fluorescence is related to a state 1 transition induced
by the oxidation of the plastoquinone pool upon illumination.
Subsequently, exposure of cells to saturating white light inten-
sities induced fluorescence quenching. A very fast quenching of
the maximal fluorescence (Fm9) was observed in wild-type cells,
especially during the first minutes of high light illumination. The
steady state (Fs) and minimal (Fo) fluorescence levels also de-
creased (Figure 2). In DOCP cells, the quenching of Fm9 was
slower than in the wild type, and no quenching of Fo was
detected (Figure 2).
Figure 1. Mutant Construction and Segregation in the Synechocystis PCC 6803 Genome.
(A) Gene arrangement of the slr1963 (encoding the OCP) and slr1964 genes. The positions of the oligonucleotides used for amplification and the
restriction sites are indicated. In the DOCP strain, the slr1963 gene was disrupted by insertion of the spectinomycin and streptomycin (Sp/Sm)
resistance cassette. In the Dslr1964 strain, the Sp/Sm resistance cassette interrupted slr1964. In the OCP-GFP mutant, the GFP gene was fused to the
C terminus of the OCP and the slr1964 gene was disrupted by the Sp/Sm resistance gene.
(B) Amplification of genomic Synechocystis DNA from the OCP-GFP mutant (lane 1), the wild type (lane 2), the DOCP mutant (lane 3), and the DSlr1964
mutant (lane 4) using car1 and car6 as primers. Lanes 5 and 6 show the PCR fragments obtained by amplification of slr1963 from the DSlr1964 strain
(lane 5) and from the DOCP mutant (lane 6) with car1 and car2 as primers. MW, 1-kb DNA ladder.
(C) Digestion of the 2- and 4-kb PCR fragments obtained using wild-type (lane 1), DOCP (lane 2), and DSlr1964 (lane 3) DNA as templates by the
restriction enzyme HincII.
994 The Plant Cell
When dark-adapted cells were exposed directly to high inten-
sities of white light, a slight increase of Fm9 was initially detected,
probably attributable to oxidation of the plastoquinone pool and
partial state 1 transition, followed by fluorescence quenching
(Figures 2B and 2C). The quenching generated in DOCP cells
was not reversible in darkness (Figure 2C). By contrast, the large
Fm9 and Fo decrease observed in wild-type cells was almost
completely reversible in the dark in the presence of chloram-
phenicol, a protein synthesis inhibitor; this finding strongly
suggested that the observed quenching was not related to D1
protein damage, which is associated with photoinhibition (Figure
2B). These results implied that, in wild-type cells, an induction of
a reversible antenna-associated NPQ occurs in high intensities
of white light that could protect PSII from photoinhibition. By
contrast, in DOCP, the decrease of maximal fluorescence ap-
peared to correlate with photoinhibition and not with NPQ.
Indeed, when the sensitivity of DOCP cells to high light
intensities was tested by following the decrease of oxygen
evolution during saturating illumination, oxygen evolution activity
decreased faster in the mutant than in the wild type, indicating
that DOCP was more sensitive to high intensities of white light
(Figure 3A). Moreover, comparison of light saturation curves of
PSII activity strongly suggested that the effective PSII antenna
size was smaller in cells that were in the quenched state. Figure
3B shows light saturation curves for oxygen-evolving activity in
wild-type cells and in 10-min photoinhibited cells. Light satura-
tion occurred at lower light intensities in wild-type control cells
than in photoinhibited cells.
Blue-Green Light–Induced Quenching and the OCP
The unexpected fluorescence characteristics of the DOCP mu-
tant prompted us to investigate the possible relationship be-
tween the OCP and the blue-green light–induced NPQ reported
previously (El Bissati et al., 2000). Blue-green light, depending on
its intensity, may be involved in state transitions or in NPQ (El
Bissati et al., 2000). Illumination of dark-adapted wild-type and
DOCP cells by dim blue-green light (which preferentially excites
PSI) increased fluorescence levels, indicating a transition to state
I induced by the oxidation of the plastoquinone pool (Figure 4).
Subsequently, exposure of dim light–adapted cells to high blue-
green light intensities induced a quenching of all fluorescence
levels in the wild type but not in DOCP (Figures 4A and 4B), even
under very high light intensities. Similar characterization of the
mutant in which the slr1964 gene, located immediately down-
stream of the ocp gene, was disrupted (DSlr1964) confirmed that
the inhibition of NPQ is attributable to the absence of the OCP
and not to the lack of transcription of slr1964 or to other polar
effects of the mutation (Figure 4C).
Transition to state II was not affected in DOCP. Illumination by
orange light (which preferentially excites PSII) of wild-type andFigure 2. Changes in Fluorescence Levels Induced by Different Inten-
sities of White Light Measured in a PAM Fluorometer.
(A) Dark-adapted wild-type and DOCP cells (3 mg chlorophyll/mL) were
successively illuminated with white light at 50 mmol�m�2�s�1 followed by
white light at 1500 mmol�m�2�s�1. Saturating pulses (1-s duration, 2000
mmol�m�2�s�1) were applied to measure Fm and Fm9 levels in darkness
and in dim light, respectively. Under high-intensity light illumination, all
centers were closed (Fs ¼ Fm9) and saturating flashes were not applied.
(B) and (C) Dark-adapted cells of the wild type (B) and DOCP (C)
were illuminated directly with high white light (1500 mmol�m�2�s�1 ) for
2 min and then incubated in darkness. During dark incubation, sat-
urating pulses were applied to measure Fm levels. Chloramphenicol was
present during all experiments.
Carotenoids and NPQ in Cyanobacteria 995
DOCP cells in state I (adapted to low intensities of blue-green light)
induced a decrease of the Fm9 level, characteristic of state II
transition, in both strains (Figures 4D and 4E). Collectively, these
data demonstrate that absence of the OCP inhibits the blue-green
light–induced NPQ and does not affect state transitions.
Is the OCP-Associated NPQ Related to Phycobilisomes or
to Chlorophyll Antennae?
In cyanobacteria, fluorescence detected at room temperature is
emitted from both chlorophyll and phycobiliproteins (Campbell
et al., 1998). In a PAM fluorometer, the measuring light has a
maximum excitation at 650 nm and the fluorescence is detected
at wavelengths of >700 nm. In cyanobacteria, which lack chlo-
rophyll b, most of the measuring light is absorbed by the
phycobilisome. This is confirmed by the low levels of fluores-
cence emitted by mutants lacking phycobilisome (El Bissati and
Kirilovsky, 2001). Thus, a decrease in the fluorescence levels
observed in a PAM fluorometer could be the result of a diminution
of the phycobilisome emission or of the chlorophyll antenna emis-
sion or of a decrease in energy transfer from the phycobilisome to
PSII. It is important to distinguish between these different possible
sources of OCP-dependent NPQ. Fluorescence spectra strongly
suggested that OCP was involved in a phycobilisome-associated
NPQ (Figure 5). At room temperature, when wild-type cells exhibit-
ing blue-green light NPQ were excited at 600 nm (light principally
absorbed by the phycobilisome), the fluorescence band with a
maximum at 660 nm (phycobilisome-related) was smaller than that
of dark-adapted wild-type control cells (Figure 5A). By contrast, the
fluorescence emission spectra were identical in DOCP cells adap-
ted either to darkness or to high intensities of blue-green light
(Figure 5C). However, when wild-type andDOCP cells were excited
by 430-nm light (principally absorbed by chlorophyll), the emission
band with a maximum at 685 nm (chlorophyll-related) increased in
cells in the quenched state compared with that in dark-adapted
control cells (Figures 5B and 5D). This chlorophyll fluorescence
increase is associated with a state II–to–state I transition upon
illumination of dark-adapted cells. Figures 5E and 5F show the 77K
fluorescence emission spectra of wild-type cells illuminated for
5 min with low (control state) or high (quenched state) intensities of
blue light. In the fluorescence spectrum generated by 600-nm
excitation, emission bands related to phycocyanin (650 nm),
allophycocyanin (660 nm), the phycobilisome terminal emitter
(685 nm), PSII (685 and 695 nm), and PSI (725 nm) were observed.
The intensity of each of these bands decreased in wild-type cells in
the quenched state (Figure 5E). By contrast, the fluorescence
spectrum generated in the quenched cells by excitation at 430 nm,
in which only PSII- and PSI-related emissions were observed, was
identical to that generated in control cells (Figure 5F). Thus, the
observed fluorescence decrease observed in the PAM fluorometer
was attributable to the quenching of the phycobilisome fluores-
cence emission and a concomitant decrease in the energy transfer
from the phycobilisome to the photosystems.
OCP-Associated NPQ in Phycobilisome and
PSII-Deficient Mutants
To substantiate the hypothesis that the OCP is specifically
involved in a phycobilisome-dependent NPQ mechanism, addi-
tional mutants were characterized: (1) lacking phycocyanin (CK,
Dcpc operon) (B. Ughy and G. Ajlani, unpublished data); (2)
lacking phycocyanin and OCP (CK-DOCP) (this work); (3) lacking
allophycocyanin (DAB, DapcAB) (Ajlani et al., 1995); (4) lacking
phycobilisome (PAL, DapcAB, DapcE, PC�) (Ajlani and Vernotte,
1998); (5) lacking PSII but retaining intact phycobilisome (DCP47
[this work] and DCP43-DpsbD; psbDI/C/DII� [Yu and Vermaas,
1990]); (6) lacking both the OCP and PSII (DCP47-DOCP) (this
work); and (7) lacking IsiA (DIsiA) (this work). The construction of
the new mutants used in this study is described in Methods.
Figure 3. Photoinhibition: Effect of High Intensities of White Light on
Oxygen Evolution in Wild-Type and DOCP Cells.
(A) Decrease of oxygen evolution induced in wild-type (circles) and
DOCP (squares) cells (10 mg chlorophyll/mL) by exposure to white light
(three lamps of 1000 mmol�m�2�s�1 each). Error bars represent SE from
three independent experiments. One hundred percent of oxygen-
evolving activity was 207 6 5 mmol O2�h�1�mg chlorophyll�1 in the wild
type and 213 6 6 mmol O2�h�1�mg chlorophyll�1 in the DOCP mutant.
(B) Saturation light curves of oxygen evolution activity in control wild-
type cells (closed circles) and in 10-min photoinhibited wild-type cells
(open circles). The photoinhibited cells were incubated on ice until
measurement. Error bars represent SE from four independent experi-
ments. One hundred percent of the oxygen-evolving activity was 205 6
5 mmol O2�h�1�mg chlorophyll�1 in control cells and 160 6 5 mmol
O2�h�1�mg chlorophyll�1 in photoinhibited cells.
996 The Plant Cell
In CK (lacking phycocyanin), the PSII antenna is composed of
the phycobilisome core containing allophycocyanin. In this mu-
tant, blue-green light still induced a reversible fluorescence
decrease (Figure 6A). The NPQ was smaller and the recovery
faster than in the wild type. This slight fluorescence quenching
was completely inhibited in the absence of the OCP (Figure 6B).
In PAL, devoid of phycobiliproteins, and in DAB, lacking allophy-
cocyanin, high intensities of blue-green light were unable to
induce fluorescence quenching (Figures 6C and 6D). By contrast,
in PAL (El Bissati and Kirilovsky, 2001) and in DAB (G. Ajlani and
C. Vernotte, unpublished data), changes in the redox state of the
plastoquinone pool induced state transitions. These results
suggest that the OCP may interact with the core of the phyco-
bilisomes to induce fluorescence quenching and support a role
for the OCP in a phycobilisome-associated NPQ.
In the DCP47 mutant, which lacks PSII reaction centers and
contains only traces of unconnected CP43 (Eaton-Rye and
Vermaas, 1991), fluorescence at room temperature is almost
entirely emitted by the phycobilisomes. As expected, because
of the lack of active PSII, no variable fluorescence was detected
in this mutant. In dark-adapted DCP47 cells, blue-green light
induced a large amount of fluorescence quenching that recov-
ered in the dark in the presence of a protein synthesis inhibitor
(Figures 7A and 7B). The quenching of the 660-nm emission
fluorescence band was even larger than that observed in wild-
type cells (Figure 7A).
The fluorescence quenching increased with blue-green light
intensity in wild-type and DCP47 cells (Figures 7C and 7D). Blue-
green light up to 160mmol�m�2�s�1 induced the transition to state
I in dark-adapted, low-fluorescence wild-type cells (data not
shown) and did not induce any quenching in the DCP47 mutant
(Figure 7D). Above this intensity, fluorescence quenching was
induced, and it increased with light intensity (Figures 7C and 7D).
Green light also induced fluorescence quenching with high
efficiency in dark-adapted and low-light-adapted cells (Figures
7C, 7E, and 7F). By contrast, orange-red light did not induce any
NPQ (Figures 7E and 7F). Similar results were obtained with the
DCP43-DPsbD mutant, lacking both interior chlorophyll anten-
nas (CP43 and CP47) and PSII (Yu and Vermaas, 1990) (data not
shown). In the DCP47-DOCP double mutant, lacking both the
OCP and the PSII core, even very high intensities of blue-green
light were unable to induce fluorescence quenching (Figure 7D).
We also examined the relationship between the OCP-associated
NPQ and the NPQ mediated by the IsiA chlorophyll binding
Figure 4. Effect of the OCP on the Development of Blue-Green Light–
Induced NPQ and on State Transitions.
(A) to (C) Measurements of fluorescence yield by a PAM fluorometer in
dark-adapted wild-type (A), DOCP (B), and DSlr1964 (C) cells illuminated
successively with low-intensity blue-green light (400 to 550 nm,
80 mmol�m�2�s�1 ) and high-intensity blue-green light (740 mmol�m�2�s�1).
(D) and (E) Measurements of fluorescence yield by a PAM fluorometer in
dark-adapted wild-type (D) and DOCP (E) cells illuminated successively
with low-intensity blue-green light (400 to 550 nm, 80 mmol�m�2�s�1) and
orange light (600 to 650 nm, 20 mmol�m�2�s�1). Blue-green illumination
induced the state 1 transition (high fluorescence state), and then orange
illumination induced the state 2 transition (low fluorescence state).
Saturating pulses separated by 30 s were applied to assess Fm9.
Carotenoids and NPQ in Cyanobacteria 997
Figure 5. Involvement of Phycobilisomes in Thermal Energy Dissipation.
(A) to (D) Room temperature fluorescence spectra of dark-adapted wild-type and DOCP cells (solid lines) and after 5 min of high-intensity blue-green
light illumination (dashed lines).
(E) and (F) 77K fluorescence spectra of wild-type cells after 5 min of illumination with low-intensity (gray lines) and high-intensity (dashed lines) blue-
green light. These spectra were normalized to fluorescence emitted by known concentrations of phycoerythrin (PE; excitation, 600 nm) or fluorescein
(excitation, 430 nm) added to the samples just before recording the spectra. APC, allophycocyanin; PC, phycocyanin.
Each spectrum shown is the mean of 12 spectra from three independent experiments (mean of four spectra per experiment). Excitation was performed
at 600 nm ([A], [C], and [E]) and at 430 nm ([B], [D], and [F]).
998 The Plant Cell
protein. Figure 7H shows that in a mutant lacking the IsiA protein,
blue-green light induced NPQ similar to that induced in the wild
type, demonstrating that this protein is not involved in the OCP-
mediated process.
Thus, the blue-green light–induced NPQ associated with OCP
can occur in the absence of the chlorophyll core antenna (CP43,
CP47, and IsiA) as well as in the absence of reaction center II.
Moreover, the kinetics of Fm9 quenching induced by strong blue-
green light were similar in the absence or presence of DCMU,
a PSII electron transport inhibitor, or nigericin, an uncoupler
(Figure 7G). These chemicals had no observable effect on the
recovery kinetics (Figure 7G). This finding indicates that the
cyanobacterial NPQ induced by blue-green light was not related
to the redox state of the plastoquinone pool or to DpH. Instead, it
seems to be related to the energy collected by the OCP, which
absorbs in the blue and green regions of the visible spectrum but
not in the orange-red region (Kerfeld, 2004b; Polivka et al., 2005).
The OCP Is Associated with Thylakoids and Phycobilisomes
The quenching phenomenon we observed suggests that the
OCP is associated with phycobilisomes and/or thylakoids. To
identify its cellular location, a Synechocystis PCC 6803 strain
containing an OCP-GFP fusion protein was prepared (Figure 1).
Panel 1 in Figure 8A shows that in the OCP-GFP mutant, green
fluorescence appeared to be distributed throughout the cell. It
is not at the periphery, indicating that OCP-GFP does not pre-
ferentially accumulate in the cell wall or in the cytoplasmic
membrane. No significant green fluorescence was observed in
wild-type cells (Figure 8A, row 2).
Cells containing the OCP-GFP fusion protein were also studied
by immunogold labeling with analysis by electron microscopy.
Visualization of the anti-GFP antibodies showed that most of the
OCP-GFP fusion proteins were localized in the interthylakoid
cytoplasmic region, the phycobilisome side of the membranes
(Figures 8B and 8C, panel 1). The density of the gold particles in
the cytoplasmic interthylakoid region (which represents 65% of
the cellular surface) was significantly higher (average ¼ 46.33 6
3.93 particles/mm2) than in the rest of the cytoplasm (average ¼17.06 6 1.78 particles/mm2; P < 10�6). Thus, 82% of the gold
particles were in the thylakoid region. In the absence of the
primary antibody, no labeling was observed (Figure 8C, panel 2).
Evidence for the interaction between the OCP and the thyla-
koids was corroborated by the presence of the OCP-GFP fusion
protein in phycobilisome-associated (MP) and phycobilisome-
free (M) membrane preparations (Figure 9). The membrane
fractions were obtained by centrifugation of cells broken in a
phosphate/citrate buffer (to obtain MP) or in a MES buffer (to
obtain M). The GFP fluorescence emission spectra of different
Figure 6. Strong Blue-Green Light Effect in Different Phycobilisome
Mutants.
Measurements of fluorescence yield by a PAM fluorometer in dark-
adapted CK ([A]; phycocyanin-deficient mutant), CK-DOCP (B), DAB ([C];
allophycocyanin-deficient mutant), and PAL ([D]; phycobilisome-deficient
mutant) cells illuminated successively with low-intensity blue-green light
(400 to 550 nm, 80 mmol�m�2�s�1; for PAL, 300 mmol�m�2�s�1) and high-
intensity blue-green light (740 mmol�m�2�s�1; for PAL, 1700 mmol�m�2�s�1).
Carotenoids and NPQ in Cyanobacteria 999
Figure 7. Blue-Green Light–Induced OCP-Related Fluorescence Quenching in DCP47 and DIsiA Mutant Cells.
(A) Room temperature fluorescence spectra of control (solid line) and quenched (dashed line) DCP47 cells. Excitation was performed at 600 nm.
(B) Fluorescence level changes in dark-adapted cells of the DCP47 mutant during successive illumination by high blue-green light followed by dark
incubation for fluorescence recovery.
1000 The Plant Cell
fractions are shown in Figures 9A and 9B. When the soluble
proteins were separated from the membrane fractions by cen-
trifugation, most of the OCP-GFP coprecipitated with the mem-
brane fractions (Figure 9A). The GFP fluorescence emission in
the M fraction was slightly smaller than that in the MP fraction,
whereas in both supernatants, the GFP emission was very small,
being smaller in the phosphate/citrate than in the MES superna-
tant. Isolation of the MP and M fractions in a sucrose gradient
gave similar results (data not shown). When the MP fraction was
resuspended in a low-phosphate buffer to dissociate the phy-
cobilisomes from the membrane, most of the OCP-GFP remained
attached to the membrane (Figure 9B). By contrast, allophyco-
cyanin and phycocyanin emissions were prominent in the MP
fraction but small in the M fraction (data not shown). These
results were supported by protein separation by SDS-PAGE and
detection of the OCP-GFP fusion protein by an anti-GFP anti-
body. This antibody reacted with a 65-kD polypeptide corre-
sponding to the expected molecular mass of the whole fusion
protein. The antibody binding was greater in the MP and M
fractions (Figure 9D) than in the supernatant fractions (Figure 9C).
Thus, our preliminary localization results suggest that there is a
relatively strong interaction between the OCP and the thylakoids,
with almost all of the OCP-GFP in the MP fraction.
The OCP was very sensitive to a membrane protease activated
during membrane solubilization (20 min, 4 or 408C) in the various
sample buffers used for protein separation by gel electrophore-
sis. Typically, we observed a band of 50 kD instead of the ex-
pected 65 kD, which corresponds to the whole fusion protein
(Figure 9D). Only solubilization of the MP and M fractions at 958C
for 3 min prevented the proteolysis (no trace of the 50-kD band
was detected). By contrast, the OCP-GFP detected in the soluble
protein fraction was not proteolyzed under any conditions. As
noted above, a 16-kD RCP corresponding to the N terminus of the
OCP was isolated from cells concomitantly with OCP. The 50-kD
band may correspond to the GFP fused to the C-terminal domain
of proteolyzed OCP. The biological significance of this derivative
form, if any, is unknown, but in several cyanobacterial genomes
(Gloeobacter violaceus, Nostoc punctiforme, and Nostoc PCC
7120), in addition to the full-length OCP gene there are other genes
corresponding to RCP-like proteins (the N-terminal domain of the
OCP) (Kerfeld, 2004b). Moreover, in Thermosynechoccocus elon-
gatus, there are single-copy open reading frames for the N- and
C-terminal domains of the OCP. These data suggest that the N-
terminal RCP may have a distinct function and/or that different
combinations of N- and C-terminal domains result in different
OCPs that may be tailored for different (photoprotective) roles.
DISCUSSION
OCP Provides Photoprotection through NPQ
The results presented in this work demonstrate that the OCP is
specifically involved in NPQ that appears to be associated with a
photoprotective energy-dissipation mechanism. In the absence
of the OCP, the NPQ induced by strong white or blue-green light
in Synechocystis PCC 6803 cells is completely inhibited, and the
cells are more sensitive to high light intensities. The observation
that the effective antenna size was smaller in the cells in the
quenched state and that the oxygen-evolving activity decreased
faster in the DOCP mutant under high-light conditions strongly
supports the hypothesis that the OCP-related mechanism dis-
sipates the excess absorbed energy, thereby decreasing the
amount of energy arriving at the photochemical centers.
OCP-Related NPQ Is Associated with Phycobilisomes
Intense blue-green light was able to induce NPQ in Synecho-
cystis PCC 6803 mutants lacking the inner chlorophyll antenna,
CP43 and CP47, but it was unable to induce NPQ in mutants
lacking phycobilisomes or the phycobilisome core. In addition,
fluorescence spectra showed that when the cells were in the
quenched state, the fluorescence emitted by the phycobilisomes
decreased and that there was less energy transfer from the
phycobilisomes to PSII and PSI. These results demonstrated that
NPQ induced by blue-green light and inhibited by the absence
of the OCP is related to the phycobilisomes. The evidence for
Figure 7. (continued).
(C) Wild-type cells adapted to low blue-green light intensities (high fluorescence state) were illuminated with strong blue-green light (400 to 550 nm) at
300 (circles), 470 (squares), and 730 (diamonds) mmol�m�2�s�1 or with green light (500 to 550 nm, 510 mmol�m�2�s�1; triangles) to induce the quenched
state.
(D) Fluorescence level changes in dark-adapted DCP47 mutant cells during illumination at 150 (dashed line), 350 (dotted line), and 1000 (solid line)
mmol�m�2�s�1 blue-green light and in dark-adapted DCP47-DOCP mutant cells during illumination at 1000 mmol�m�2�s�1 blue-green light (solid line).
The DCP47 and DCP47-DOCP mutants lack variable fluorescence because of the absence of PSII reaction centers.
(E) Dark-adapted wild-type cells were illuminated with orange-red (600 to 650 nm; squares), blue-green (400 to 550 nm; circles), or green (triangles) light
at 470 mmol�m�2�s�1. Saturating pulses separated by 30 s were applied to assess Fm9. The orange-red light at this intensity closed 95% of PSII centers,
whereas blue-green and green light closed only 20 and 10% of centers, respectively.
(F) Fluorescence level changes in dark-adapted DCP47 cells illuminated by orange-red (600 to 700 nm, 3000 mmol�m�2�s�1; dashed line), green (500 to
550 nm, 470 mmol�m�2�s�1; dotted line), and blue-green (400 to 550 nm, 470 mmol�m�2�s�1; solid line) light.
(G) Measurements of fluorescence yield by a PAM fluorometer for wild-type cells adapted to low blue-green light intensities (high fluorescence state) in
the presence of DCMU (solid line), nigericin (squares), or without additions (circles) illuminated for 200 s with strong blue-green light (740 mmol�m�2�s�1 )
to induce the quenched state and then illuminated with low blue-green light (80 mmol�m�2�s�1) to allow fluorescence recovery.
(H) Measurements of fluorescence yield by a PAM fluorometer for dark-adapted DIsiA cells illuminated successively with low-intensity blue-green light
(400 to 550 nm, 80 mmol�m�2�s�1) and high-intensity blue-green light (740 mmol�m�2�s�1).
Carotenoids and NPQ in Cyanobacteria 1001
an interaction between the OCP and the phycobilisomes and
thylakoids necessary for this NPQ was supported by the coiso-
lation of the OCP-GFP fusion protein with the phycobilisome-
associated membrane fraction. Our results are in accord with
those of a proteomic study of Synechocystis PCC 6803 thyla-
koids (Srivastava et al., 2005) in which the OCP was observed in
a purified thylakoid preparation. Furthermore, our studies by im-
munogold labeling and electron microscopy showed that most
of the OCP-GFP fusion protein is present in the interthylakoid
region of the cell.
This NPQ also occurs in a Synechocystis PCC 6803 mutant
lacking phycocyanin (CK) in which only phycobilisome cores are
present. This finding suggests that the OCP interacts with a
component of the phycobilisome core. The Lcm, the linker-
membrane protein that acts as the terminal energy acceptor in
the phycobilisome, is a good candidate for this role. The faster
recovery from the quenched state in CK compared with the wild
type implies a weaker interaction between the OCP and the
phycobilisome and/or the involvement of the movement of
the phycobilisome relative to the membrane, as suggested by
Joshua et al. (2005).
Under Fe-starvation conditions, a large reversible quenching
of Fo and Fm levels, suggested to be mediated by the IsiA
protein, is also induced by blue light (Cadoret et al., 2004; Bailey
et al., 2005; Joshua et al., 2005). Because we demonstrated that
the blue-green light–induced NPQ is not mediated by the IsiA
protein, we propose that the blue light–induced NPQ observed
under Fe-starvation conditions is also related to phycobilisome
and OCP; the presence of IsiA only increases the extent of the
quenching. If part of the phycobilisomes transfer absorbed en-
ergy to IsiA complexes, that contributes to the Fo level (Joshua
et al., 2005); in cells containing quenched phycobilisomes, less
energy will arrive at the IsiA complexes and a larger fluorescence
quenching will be observed. This blue light mechanism seems
to be independent of the quenched state of the large IsiA
oligomers that appear in long-term iron-depleted cells, as
Figure 8. In Situ Localization of the OCP-GFP Fusion Protein.
(A) Distribution of green fluorescence. Phase contrast (left), GFP fluorescence (middle), and chlorophyll and phycobiliprotein fluorescence (right)
micrographs of OCP-GFP (row 1) and wild-type (row 2) cells are shown. Bar ¼ 3 mm.
(B) Histogram of gold particle density in whole cells (bar 1), interthylakoid region (bar 2), and non-thylakoid-related cytoplasm (bar 3). Thirty-three cells
were analyzed and 1120 gold particles were counted. On average, the thylakoid region represents 65% of the cellular surface. Error bars indicate SE.
(C) Immunogold labeling of a thin section of OCP-GFP–transformed cells. Panel 1, OCP-GFP cells were labeled with a polyclonal antibody against the
GFP coupled to 10-nm gold particles; panel 2, no labeling was observed without the primary antibody. Bar ¼ 0.5 mm.
1002 The Plant Cell
already suggested by Ihalainen et al. (2005). Energy dissipation in
IsiA aggregates was independent of the quality of the excitation
light.
NPQ Appears to Be Induced via Activation of the OCP by
Light Absorption
In higher plants, the antenna-associated NPQ is induced by low
pH in the thylakoid lumen. By contrast, the phycobilisome-
associated NPQ described here is not dependent on the pres-
ence of a transthylakoid DpH, as shown by the inability of
uncouplers to affect the kinetics of quenching and recovery.
Several lines of evidence indicate that the induction of the
quenching is also independent of excitation pressure on PSII or
changes in the redox state of the plastoquinone pool. On the one
hand, the intensities of blue light that induce the NPQ are not
saturating for oxygen evolution, and NPQ can occur when a small
number of PSII centers are closed (El Bissati et al., 2000). On the
other hand, NPQ could be induced under conditions in which the
plastoquinone pool becomes (1) more reduced (by transfer from
darkness or dim blue light to high intensities of white light) or (2)
more oxidized (when going from dark to blue light, in the pres-
ence of DCMU) or (3) when its redox state is unaffected (in the
PSII-lacking mutant). In addition, because blue light is absorbed
by PSI, we cannot discount the possible involvement of PSI
activity, but this is unlikely because green light is very efficient in
the induction of NPQ.
Recently, two independent studies showed that in two different
Synechocystis PCC 6803 mutants lacking PSII, blue-green light
induces a quenching of phycobilisome emission (Rakhimberdieva
et al., 2004; Scott et al., 2005). The action spectrum for the
Figure 9. GFP Fluorescence Measurement, Gel Electrophoresis, and
Protein Gel Blot Analysis of Soluble and Membrane Fractions.
(A) Fluorescence emission spectra. GFP emission in the membrane frac-
tions MP (for membrane-bound phycobilisomes) and M (for membrane-free
phycobilisomes) at 4 mg chlorophyll/mL and the soluble fractions sup MES
(for MES supernatant) and sup P/C (for phosphate-citrate supernatant) at
a concentration corresponding to that of the membrane fractions (see
Methods). Excitation was at 480 nm. Values shown are means of four
independent experiments.
(B) Fluorescence emission spectra. GFP in the MP fraction (3.5 mg
chlorophyll/mL) and in the M and phycobilisome (PBS) fractions obtained
after suspension of the MP fraction in MES buffer (1-h incubation) and
subsequent centrifugation. Values shown are means of three different
experiments.
(C) Coomassie blue–stained gel electrophoresis and immunoblot detec-
tion of the OCP-GFP fusion protein of the M fraction (lane 1), the MP
fraction (lane 2), and the soluble fractions of cells broken in P/C buffer
(lane 4) and MES (lane 5); lane 3, molecular mass markers. In the
immunoblot (lane 3), the heavy chain of the mouse IgG is visualized.
(D) Coomassie blue–stained gel electrophoresis and immunoblot detec-
tion of the OCP-GFP fusion protein of the MP (lanes 2 and 4) and M (lanes
1 and 5) fractions solubilized for 3 min at 958C (lanes 1 and 2) or for 20 min
at 48C (lanes 4 and 5); lane 3, molecular mass markers. In the immunoblot
(lane 3), the heavy chain of the mouse IgG is visualized.
Each slot contained 2 mg of chlorophyll of the MP or M fraction or the
corresponding volumes of the supernatants (see Methods). The rela-
tionship between the volumes of membrane and soluble fractions was
the same for the fluorescence spectra and gels.
Carotenoids and NPQ in Cyanobacteria 1003
phycobilisome quenching, in both cases, was reminiscent of a
carotenoid absorption spectrum. Moreover, Rakhimberdieva
et al. (2004) proposed that a carotenoid activated by blue light
induces the reversible NPQ. Here, we have shown that the ab-
sence of the OCP completely inhibits the blue light–induced
phycobilisome-related NPQ in PSII-lacking mutants and in the
wild type, demonstrating that this carotenoid is associated with
the OCP. The action spectra (Rakhimberdieva et al., 2004)
resembled the absorption spectra of the OCP in Arthrospira
maxima (Polivka et al., 2005). We cannot completely rule out the
involvement of a cryptochrome or a BLUF protein as the blue light
sensor, but the fact that green light (500 to 550 nm) induced NPQ
very efficiently renders this improbable. The oxidized flavin ade-
nine dinucleotide, the cofactor of these blue photoreceptors, has
a similar spectrum in the blue region, but it absorbs very weakly in
the green region (for the BLUF [Slr1694] spectrum of Synecho-
cystis PCC 6803, see Masuda et al., 2004).
Working Models for OCP Activity
Many questions arise from our findings about the role of the OCP
in the phycobilisome-related NPQ. First, what are the changes
that are induced in the carotenoid and in the protein by high light
intensities that activate the OCP to induce the quenching? For
example, blue-green light absorption could lead to carotenoid
isomerization, inducing conformational changes in the chromo-
phore and in the protein. Alternatively (or in addition), proton or
electron transfer could be induced as in the flavin-containing
blue receptors. Changes in the tertiary or quaternary structure
(OCP is a dimer) or proteolysis of the protein may also be involved
in the mechanism.
Our results do not permit conclusions to be drawn on the
mechanism by which energy is dissipated. There are several
possibilities. The OCP could be the light sensor, a mediator of
quenching, the quencher itself, or play a combination of these
roles. The activated OCP, through interaction with the core of the
phycobilisome, could cause an alteration of the phycobilisome
structure, leading to the quenched state. Alternatively, the ca-
rotenoid of the OCP could interact directly with a phycobilin
chromophore (most probably the terminal acceptor) and dissi-
pate the absorbed energy.
This study demonstrated that the OCP is specifically involved
in a phycobilisome-associated NPQ and not in other mecha-
nisms affecting the levels of fluorescence (e.g., state transitions
or D1 damage). This mechanism, induced by both blue-green
light and saturating intensities of white light, likely protects PSII
from high light exposure. We report here a novel function for a
soluble carotenoid protein as a mediator of a photoprotective
response. Our results reveal the role of the OCP and confirm the
hypothesis that in cyanobacteria, as in higher plants, there exists
a mechanism involving energy dissipation in the antenna (phy-
cobilisomes). The molecular mechanism of this novel process
awaits elucidation. We propose a working model in which the
OCP acts as a photoreceptor that responds to blue-green light
and subsequently induces energy dissipation (and fluorescence
quenching) through interaction with the phycobilisome core.
Further testing of this hypothesis is likely to open new frontiers in
carotenoid function.
METHODS
Plasmid Construction
DOCP Plasmid
The ocp gene (slr1963) was amplified by PCR using genomic DNA of
Synechocystis PCC 6803 as template. Two synthesized oligonucleotides
were used as primers: car1, 59-CTACGCGGATCCATTGACTCTGCCCG-
CGGAATTT-39, and car2, 59-CGGCCGCTCGAGCAAAGTTGAGTAATT-
CTTTGGG-39. The resulting PCR product was digested by EagI and AvaI
restriction enzymes and then cloned in the polylinker EagI-XhoI restriction
sites of pBluescript SKþ (Stratagene) plasmid. The slr1963 gene was
interrupted by inserting a 2.2-kb DNA fragment (v cassette) containing
the aadA gene from Tn7, conferring resistance to Sp/Sm, in the unique
restriction site HincII.
OCP-GFP Plasmid
The slr1963gene ofSynechocystisPCC 6803 was amplified by PCR using
the oligonucleotides car1 and car4 (59-CTGAAGGGAGTTAGGATCCC-
GAGCAAAGTTGAG-39) containing a BamHI restriction site. The stop
codon was suppressed. The PCR product was cleaved with SphI and
BamHI restriction enzymes and cloned into the polylinker SphI and
BamHI restriction sites of a pEGFP ampicillin-resistant vector to create a
recombinant slr1963-GFP gene. In addition, the immediately downstream
slr1964 open reading frame, encoding a hypothetical protein, was am-
plified by PCR using two other oligonucleotides: the NotI-creating primer
(car5,59-TTACTAACTTTGGCGGCCGCAATAACTCCCTTCAGAG-39)and
the SpeI-creating primer (car6, 59-CACCGGACTAGTCAAAAACTAT-
CTGCTGGCGATCG-39). This second PCR fragment was cloned into
the pEGFP/OCP ampicillin-resistant vector opened with the same en-
zymes. The slr1964 gene was inactivated by insertion of the Sp/Sm
cassette in the unique ClaI restriction site.
Dslr1964 Plasmid
The OCP-GFP plasmid was cleaved with both NotI and SpeI restriction
enzymes. The digestion product of 3.1 kb contained the slr1964 gene
inactivated by insertion of the Sp/Sm cassette in the unique ClaI restric-
tion site. This DNA fragment was cloned into a pBluescript SKþ ampi-
cillin-resistant vector opened with the same enzymes to obtain the
Dslr1964 plasmid.
DisiA Plasmid
The isiA gene (sll0247) of Synechocystis PCC 6803 was amplified by PCR
and cloned in the polylinker SrfI restriction site of a pPCR-Script SKþampicillin-resistant vector. The isiA gene was inactivated by inserting
Sp/Sm cassette in the unique PmlI restriction site.
DCP47 Plasmid
The DNA region of Synechocystis PCC 6803 containing the psbB gene,
encoding the CP47 protein, was amplified by PCR using the oligonucle-
otides BM (59-CATGGTGATAATCAAGGGATG-39) and BK (59-CGCTT-
TCGTCGTGGCCGGTAC-39). The PCR fragment containing the psbB
gene was cloned onto the EcoRV site of the pBC SKþ plasmid. In the
resulting plasmid, a 550-bp BstEII fragment was substituted by the eryth-
romycin resistance cassette.
Transformation, Selection, and Genetic Analysis of Mutants
The DOCP, OCP-GFP, DisiA, DCP47, and Dslr1964 plasmid constructs
were used to transform wild-type Synechocystis sp PCC 6803. To obtain
1004 The Plant Cell
the double mutants DCP47-DOCP and CK-DOCP, DCP47 and CK cells
were transformed with the DOCP plasmid. Transformants were selected
under dim light at 328C on plates containing different antibiotics: 25mg/mL
spectinomycin and 12 mg/mL streptomycin, 20 mg/mL erythromycin, or
40 mg/mL kanamycin. The nonphotosynthetic DpsbB mutant was grown
in the presence of 20 mM glucose because it has an obligate heterotroph
phenotype.
Genomic DNA was isolated from Synechocystis PCC 6803 essentially
as described by Cai and Wolk (1990). To confirm the homoplasmicity and
complete segregation of the different mutants, PCR analysis and specific
digestions by restriction enzymes were performed.
Culture Conditions
Wild-type and mutant cells were grown photoautotrophically in a mod-
ified BG11 medium as described by Herdman et al. (1973) containing
twice the concentration of sodium nitrate. Cells were shaken in a rotary
shaker (120 rpm) at 308C and illuminated by fluorescent white lamps
giving a total intensity of ;30 to 40 mmol�m�2�s�1 under a CO2-enriched
atmosphere. The selected mutants were grown in the presence of the
appropriate antibiotics. The cells were maintained in the logarithmic phase
of growth and were collected having optical densities of 0.6 to 0.8 at 800 nm.
Photoinhibition
For the photoinhibition experiment (Figure 3A), cells (10 mg chlorophyll/
mL) were incubated at 308C in a glass tube (3 cm diameter) under stirring
and illuminated with three Atralux spots of 150 W (1000 mmol�m�2�s�1
each lamp). The effective intensity of light absorbed by each cell was
much less as a result of self-absorption by the cells in the glass tube. For
comparison, when wild-type cells (10mg chlorophyll/mL) were illuminated
in the PAM cuvette (1500 mmol�m�2�s�1), oxygen-evolving activity was
abolished within 30 min. Therefore, we estimate the actual amount of
incident light to be <1500 mmol�m�2�s�1. For the experiment shown in
Figure 3B, wild-type cells were illuminated for 10 min (same conditions as
in Figure 3A) and then incubated on ice to inhibit the recovery of NPQ
(data not shown) until oxygen evolution measurements could be com-
pleted. Oxygen evolution of intact cells (10 mg chlorophyll/mL) was
measured polarographically using a Clark-type oxygen electrode with the
addition of 1 mM 2,6-dichlorobenzoquinone as an artificial PSII electron
acceptor.
Fluorescence Measurements and Detection
In the course of our studies, we always observed the blue-green light–
induced NPQ in wild-type cells; however, the extent of the reversible NPQ
and the kinetics of recovery were sensitive to cell concentration, phyco-
bilisome concentration, PSII:PSI and Fv:Fo ratios, etc. Thus, wild-type
and mutant cells were grown in similar conditions (see Culture Conditions)
and collected at an equal cell concentration and phycocyanin:chlorophyll
ratio. The yield of chlorophyll fluorescence was monitored in a modulated
fluorometer (PAM; Walz, Effelrich, Germany) adapted to a Hansatech
oxygen electrode as described previously (El Bissati et al., 2000). All NPQ
induction and recovery experiments were performed in a stirred cuvette
of 1 cm diameter (328C) at a chlorophyll concentration of 3 mg/mL in the
presence of chloramphenicol (30 mg/mL) to inhibit protein synthesis.
Recovery was in darkness or under 50 to 80 mmol�m�2�s�1 blue-green
light. The nomenclature used was as follows: Fo, minimal fluorescence
level, the fluorescence emitted by open reaction centers in dark-adapted
cells; Fm, maximal fluorescence level in dark-adapted samples; Fm9,
maximum fluorescence under illumination, corresponding to the fluores-
cence emitted by the maximum concentration of closed reaction centers;
Fs, steady state fluorescence level; Fv, variable fluorescence ¼ Fm � Fo.
The Fo level was determined by illuminating dark-adapted cells with a low
intensity of red-modulated light (pulses of 1 ms, 1.6 kHz, 0.024 mmol�m�2�s�1). Saturating pulses (2000 mmol�m�2�s�1, 1 s) were applied to
measure Fm and Fm9 levels. Application of such pulses that transiently
close all PSII centers serves to distinguish between photochemical
quenching and NPQ.
The fluorescence emission spectra were recorded on a Hitachi F-3010
fluorescence spectrophotometer. Excitation was done at 600 or 430 nm.
The cells were at a concentration of 5mg chlorophyll/mL. Phycoerythrin or
fluorescein (1.75mM) was added to facilitate normalization of the spectra.
Phycoerythrin (isolated from Rhodella violacea) was added to obtain a
phycoerythrin emission slightly higher than those of Synechocystis phy-
cocyanin and allophycocyanin.
To detect green fluorescence from the GFP-OCP, cyanobacteria were
deposited on a glass slide and mounted for observation. Micrographs
were taken with an Olympus VAN-Ox AH2 fluorescence microscope fitted
with a color cooled charge-coupled device camera (Coolsnap; Roper
Scientific).
MP and M Membrane Preparations
Cells were resuspended in a buffer of 0.5 M K-phosphate and 0.3 M Na-
citrate (pH 6.8) (P/C buffer) to obtain MP and in a 20 mM MES, pH 6.8,
buffer to obtain M at a chlorophyll concentration of 1 mg/mL and broken
in a mini-bead-beater in the presence of glass beads. The M and MP
fractions were collected by centrifugation and frozen at –808C until use for
gel electrophoresis. The homogenates containing the broken cells were
loaded on a continuous sucrose gradient (0 to 50%) prepared in the P/C
buffer and centrifuged. The colored band containing the MP fraction or
the M fraction was collected, pelleted by centrifugation, and stored at
�808C. We are aware that the M and MP fractions obtained only by
centrifugation could be contaminated by cytoplasmic membranes. How-
ever, similar results on detection of the OCP-GFP fusion protein were
obtained using the M and MP fractions collected directly by centrifugation
and those collected by the sucrose gradient that are more purified (data
not shown).
The OCP-GFP in the M, MP, and soluble fractions compared in protein
gel blots and fluorescence spectra (Figure 9) was isolated and quantified
as follows: 0.5 mL of broken cells (1 mg chlorophyll/mL) (in MES or P/C
buffer) was centrifuged. The pellet containing the membrane fraction was
resuspended in 100mL of MES buffer. The supernatant was concentrated
to 100 mL. We assumed that 1 mL of the concentrated supernatant
corresponded to ;1 mL of the resuspended membrane fraction. This
assumption was confirmed by the observation that the chlorophyll:
phycocyanin absorption ratio in a solution containing 5 mL of supernatant
and 5 mL of membranes was equivalent to that of whole cells. If
necessary, adjustments were made by dilution to equalize the ratios
between the two samples. For the fluorescence spectra shown in Figure
9B, 20 mL of MP suspended in P/C buffer was diluted with 2 mL of MES
buffer and incubated on ice for 1 h. The membranes were subsequently
precipitated by centrifugation and resuspended in 100 mL of MES. The
supernatant was concentrated to 100 mL. Again, the chlorophyll:phyco-
cyanin ratio was used to normalize the samples.
Gel Electrophoresis and Protein Gel Blots
Proteins of the MP and M fractions and of the soluble fractions of the
OCP-GFP mutant were analyzed by SDS-PAGE on a 12% polyacryla-
mide/2 M urea gel in a Tris/MES system (Kashino et al., 2001). The OCP-
GFP fusion protein was detected by a monoclonal antibody against GFP
(Clontech).
Electron Microscopy
Samples were fixed in 4% paraformaldehyde and embedded in Unicryl.
For immunogold labeling, the sections were incubated with the primary
Carotenoids and NPQ in Cyanobacteria 1005
antibody (rabbit polyclonal antibody against GFP [Abcam]; 3 mg/mL final
dilution) in buffer T (20 mM Tris-HCl, 154 mM NaCl, 0.1% NaN3, 0.1%
BSA, 0.05% Tween 20, and 0.1% fish gelatin); then, after washing, they
were incubated with a 1:20 dilution of 10-nm gold-conjugated anti-rabbit
antibodies (Amersham) and stained in 2% uranyl acetate followed by lead
citrate. The sections were observed on a Philips EM 400 electron
microscope (FEI).
Accession Numbers
Sequence data from this article can be found in the GenBank/EMBL data
libraries under the following accession numbers: slr1963, NP_441508;
slr1964, NP_441509; isiA (sll0247), NP_441268; psbB (slr0906),
NP_442388; psbC (sll0851), NP_441119; psbD (sll0849), NP_441120;
psbD2 (slr0927), NP_442780; apcA (slr2067), NP_441194; apcB (slr1986),
NP_441195; apcA (slr2067), NP_441194; apcB (slr1986), NP_441195;
apcE (slr0335), NP_441972; cpcA (sll1578), NP_440551; cpcB (sll1577),
NP_440552; cpcC2 (sll1579), NP_440550; cpcC1 (sll1580), NP_440549;
cpcD (ssl3093), NP_440548.
ACKNOWLEDGMENTS
We thank W. Vermaas for the gift of the DpsbD-DpsbC mutant, A.
William Rutherford for stimulating discussions and critical reading of the
manuscript, Bernard Lagoutte for helpful advice and discussions, Estelle
Delphin for the low-temperature fluorescence spectra, and Krisztian Cser
for the light saturation curves of oxygen evolution. This research was
partially supported by European Union network INTRO2.
Received December 7, 2005; revised February 6, 2006; accepted Feb-
ruary 17, 2006; published March 10, 2006.
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Carotenoids and NPQ in Cyanobacteria 1007
DOI 10.1105/tpc.105.040121; originally published online March 10, 2006; 2006;18;992-1007Plant Cell
Adjélé Wilson, Ghada Ajlani, Jean-Marc Verbavatz, Imre Vass, Cheryl A. Kerfeld and Diana KirilovskyCyanobacteria
A Soluble Carotenoid Protein Involved in Phycobilisome-Related Energy Dissipation in
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