Chapter 37
Cyclic Electron Transfer Around Photosystem I
Pierre Joliot∗ and Anne Joliot
CNRS UMR 7141, Institut de Biologie Physico-Chimique, 13, rue Pierre et Marie Curie, 75005
Paris, France
Giles Johnson
University of Manchester, School of Biological Sciences, 3.614 Stopford Building, Oxford Road,
Manchester, M13 9PT, UK
Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 639
I.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640
II.
Early Observations of Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640
III. Possible Pathways of Electron Flow in Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640
IV. Redox Poising of the Cyclic Electron Transfer Chain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641
V.
Structural Organization of Thylakoid Membranes – Consequences for Cyclic Electron Transfer . . . . . . . . . . . 642
VI. Occurrence of Cyclic Flow in Higher Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643
VII. Pathway of Cyclic Flow in Higher Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647
A. NADPH-Dependent Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647
B. Ferredoxin-Dependent Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647
VIII. Cyclic Flow in Green Unicellular Algae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649
IX. Cyclic Flow in Cyanobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650
X.
Functions and Regulation of Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651
XI. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 652
Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 652
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 652
Summary
Cyclic electron transport around Photosystem I remains one of the last great enigmas in photosynthesis research.
Although first described in 1955 by Arnon and coworkers, the molecular details of the pathway, its physiological
role and even its very occurrence remain in question. Nevertheless, significant progress is starting to be made in our
understanding of this process. At least two pathways of cyclic electron transport appear to operate, one involving
the transfer of electrons from NADPH to plastoquinone and the other operating via the donation of electrons from
ferredoxin to plastoquinone. The relative importance of these two pathways seems to vary between cyanobacteria,
unicellular green algae and higher plants as do many details concerning the regulation of the pathway and its
functional organization in the thylakoid membrane. Two distinct functions for cyclic electron transport can be
defined — the generation of ATP and, in higher plants, the generation of pH to regulate light harvesting. These
two functions give rise to the need for different regulatory processes to control the ratio of cyclic and linear electron
flow. We discuss recent findings that cast new light on how cyclic electron transport is regulated under a range of
physiological conditions.
∗
Author for correspondence, email: pjoliot@ibpc.fr
John H. Golbeck (ed): Photosystem I: The Light-Driven Plastocyanin:Ferredoxin Oxidoreductase, 639–656.
C 2006 Springer.
640
I. Introduction
The concept of cyclic electron transport (ET) around
Photosystem (PS) I is sufficiently established that it features in the diagrams of photosynthetic electron transport found in every undergraduate biochemistry textbook. In spite of this celebrity, the redox components
involved, the functional importance, the regulation and,
indeed, even the very occurrence of this pathway all remain unclear. Part of the problem in studying cyclic ET
has been, that, by its very nature, it is difficult to quantify — as a cycle it involves no net flux and so we are
forced to resort to use indirect means to deduce its existence. There is now growing evidence, however, that
cyclic ET does indeed occur in a variety of organisms
under a wide range of conditions, fulfilling at least two
distinct functions. Here, we present a review of this
evidence, focusing in particular on advances over the
last decade and identifying the challenges that remain.
For more detailed coverage of earlier work and alternative views on the subject, a number of good reviews are
available (Heber and Walker, 1992; Fork and Herbert,
1993; Bendall and Manasse, 1995; Heber, 2002; Allen,
2003).
II. Early Observations of Cyclic Electron
Transfer
Photosynthetic phosphorylation (photophosphorylation) was discovered in 1954 by Arnon et al. (1954) who
established that illumination of isolated chloroplasts in
the presence of oxygen-induced ATP synthesis. As this
process was not associated with oxygen formation or
consumption, the contribution of a respiratory chain
could be excluded. Later, it was established that the dependence of photophosphorylation on oxygen can be
abolished by the addition of vitamin K or other naphthoquinones (Arnon, 1955). This experiment marks the
discovery of the cyclic phosphorylation process, (reviewed in Arnon et al., 1961). The reaction previously
characterized in 1954 by Arnon and coworkers appears
now to be a pseudocyclic process in which electrons
are transferred from water to O2 (Mehler, 1951) via the
Abbreviations: cyt – cytochrome; DBMIB – 2,5-dibromo3-methyl-6-isopropyl- p-benzoquinone; DCMU – 3-(3,4dichloro-phenyl)-1,1-dimethylurea; ET – electron transport;
ETC – electron transfer chain; Fd – ferredoxin; FNR – ferredoxin: NADP oxidoreductase; FQR – ferredoxin: plastoquinone
reductase; HQNO – 2-heptyl-4-hydroxy-quinoline N-oxide;
NDH – NADH dehydrogenase; PC – plastocyanin; PMS – Nmethylphenazonium-3-sulfonate; PQ – plastoquinone; PQH2 –
plastoquinol; PS – Photosystem; RC – reaction center.
Pierre Joliot, Anne Joliot and Giles Johnson
linear electron transfer chain. Subsequently, nonphysiological compounds, such as phenazine methosulfate
(PMS), were shown to catalyze cyclic photophosphorylation more efficiently than vitamin K (Jagendorf and
Avron, 1958).
The anaerobic cyclic phosphorylation identified in
chloroplasts appeared analogous to a cyclic phosphorylation process previously identified in chromatophores
from photosynthetic bacteria (Frenkel, 1954). In Rhodospirillum rubrum, this cyclic process was observed
to involve cytochrome (cyt) c (Smith and Baltscheffsky, 1959); in chloroplasts cyt f is involved (Arnon,
1959). In 1960, Hill and Bendall put forward a model
describing the linear electron transfer chain involving
both the photoreactions PS II and PS I working in series (Hill and Bendall, 1960). It became clear, however,
that cyclic photophosphorylation involves only PS I,
as it operates in the presence of specific inhibitors of
oxygen evolution, such as 3-(3,4-dichloro-phenyl)-1,1dimethylurea (DCMU). Tagawa et al. (1963b) established that ferredoxin (Fd) was able to catalyze cyclic
phosphorylation. This provided evidence for a physiological catalyst of cyclic ET, making it seems likely
that this process was more than an artifact of in vitro
conditions.
III. Possible Pathways of Electron Flow
in Cyclic Electron Transfer
There is a general agreement that both PS I and the cyt
b/ f complex are obligatory involved in cyclic electron flow — cyclic ET is efficiently driven by farred light and is sensitive to inhibitors of the cyt b/ f
complex, such as 2,5-dibromo-3-methyl-6-isopropylp-benzoquinone (DBMIB) and stigmatellin — however, the electron pathway from the acceptor side of PS
I back to the cyt b/ f complex has not yet been clearly
elucidated. Cyclic ET, as conventionally measured,
leads to the generation of a trans-thylakoid pH gradient. Thus, whatever the pathway involved a protonpumping step involving the cyt b/ f complex must be
postulated.
Based mainly on studies of the effects of the inhibitor antimycin A, it has been postulated that at least
two pathways exist. Experiments performed on isolated
chloroplasts showed that antimycin inhibits cyclic flow
in the presence of Fd (Tagawa et al., 1963b) but not
in the presence of vitamin K or PMS (Whatley et al.,
1959). Hosler and Yocum (1985) measured the ratio of
P/O in the presence of Fd, using oxygen as a terminal
electron acceptor and found this to be sensitive to antimycin. They explained this effect as being due to the
Chapter 37
Cyclic Electron Transfer Around Photosystem I
641
Fig. 1. Possible pathways for cyclic electron flow.
inhibition of cyclic ET. By contrast, when the same experiment was performed using NADP as an electron acceptor, the P/O ratio was high and antimycin-insensitive
(Hosler and Yocum, 1985). Scheller (1996) measured
the rereduction of P700 following light flashes in the
presence of DCMU and noted an antimycin-insensitive
portion in Fd-mediated cyclic ET, possibly suggesting a
third pathway (Scheller, 1996). However, the very slow
rate of this, barely distinguishable from the DBMIBinhibited rate, makes it hard to separate this from background redox equilibration of the sample. Cleland and
Bendall (1992) measured the oxidation of reduced Fd
following illumination of DCMU-poisoned thylakoids
and found this rate to be inhibited by antimycin A as
completely as by stigmatellin (Cleland and Bendall,
1992). One can thus conclude that several mechanisms
could be involved in a cyclic ET.
In a conventional Q-cycle process (Mitchell, 1975;
Crofts et al., 1983), electrons are transferred to the cyt
b/ f complex via a reduced quinone that binds site Qo ,
on the lumenal side of this complex. Assuming this
is the step generating pH in the cyclic process, we
need to invoke an enzyme that is able to transfer electrons from NADP or Fd to plastoquinone (PQ), with
the resultant plastoquinol (PQH2 ) being protonated
on the stromal side of the membrane. In cyanobacteria,
the presence of respiratory and photosynthetic electron
transport chains in the same membrane means that this
function could be fulfilled by a respiratory NADH dehydrogenase (NDH; complex I). Genes coding for such
an enzyme have also been identified in the chloroplast
genome of higher plants (Shinokazi et al., 1986), which
is a likely candidate to be involved in the cyclic electron
transfer chain (ETC) (Fig. 1, pathway 1).
Moss and Bendall (1984) noted that, whilst both Fdmediated and artificially mediated cyclic ET are sensitive to the cyt b inhibitor 2-heptyl-4-hydroxy-quinoline
N-oxide (HQNO), only Fd-mediated cyclic ET was
sensitive to antimycin (Moss and Bendall, 1984). This
led them to suggest an alternative site for antimycin
inhibition on an enzyme distinct from the b/ f complex, termed ferredoxin:plastoquinone oxidoreductase
(FQR) (Fig. 1, pathway 2).
The involvement of ferredoxin:NADP oxidoreductase (FNR) in cyclic ET has been much discussed. Its
involvement in the NADP-dependent pathway is presumed; however, a role in the Fd-dependent pathway
has also been postulated. The observation that this enzyme is stoichiometrically bound to the cyt b/ f complex in spinach provides an intriguing indication of an
additional role, other than that of linear electron transport (Zhang et al., 2001). It is suggested that FNR mediates the transfer of electrons from ferredoxin to site
Qi (Fig. 1, pathway 3).
The mechanisms involved in these different pathways will be discussed in more detail below.
IV. Redox Poising of the Cyclic Electron
Transfer Chain
Photochemical charge separation at the level of a
reaction center (RC) requires the presence of a reduced primary donor and an oxidized electron acceptor.
642
Over-reduction or overoxidation of the cyclic chain will
thus inhibit this process. The concept of redox poising
of the cyclic ETC was introduced to explain the oxygen
requirement of cyclic phosphorylation (Tagawa et al.,
1963a; Whatley, 1963; reviewed in Allen, 1983). Later,
Arnon and Chain (1977) established that a maximum
efficiency of the cyclic ET in the presence of oxygen
is observed in the presence of NADPH and subsaturating concentrations of DCMU that impose an optimal
redox poise of the carriers involved in the cyclic chain
(Arnon and Chain, 1977).
Redox poising is determined by the relative rate of
electron efflux or influx from or toward the carriers
belonging to the cyclic electron transfer chain. Both
efflux and influx will occur preferentially at the level
of mobile carriers that are common to linear and cyclic
chains. PS II will be the main source of reductive power
(Fig. 1, pathways a and b) while electron efflux will
occur at the level of PS I acceptors toward O2 and the
Benson–Calvin cycle (Fig. 1, pathways c and d, respectively). If we assume that cyclic and linear pathways are
connected, i.e., share the same mobile carriers, the redox poise of the cyclic chain will be controlled by the
electron flow through the linear chain. Under strong illumination, given under condition where the Benson–
Calvin cycle is inhibited (e.g., in isolated thylakoids or
dark-adapted leaves), PS II will induce an overreduction of the cyclic chain, via pathway a or b (Fig. 1).
Conversely, under conditions where PS II is inhibited
or under far-red light, the electron efflux via pathway c
or d will induce an overoxidation of the cyclic chain. On
the other hand, if the cyclic and linear chains are structurally separated, the redox poise of the cyclic chain
will be controlled by the rate of slow electron leaks
that occur between carriers involved in both processes.
We thus conclude that the structural organization of
membrane proteins that controls the localization of the
cyclic and linear chains within the membrane may play
an essential role in the control of the efficiency of the
cyclic process.
V. Structural Organization of Thylakoid
Membranes – Consequences for
Cyclic Electron Transfer
Oxygenic photosynthetic organisms — higher plants,
unicellular algae, and cyanobacteria — differ substantially in the supramolecular organization of their photosynthetic membranes and these differences may have
important consequences for the pathway and regulation
of cyclic ET.
Pierre Joliot, Anne Joliot and Giles Johnson
In higher plants, PS II and PS I are localized in different membrane regions (Andersson and Anderson,
1980). Most of PS II is localized in the appressed
regions of the grana stacks while PS I is localized
in the stroma lamellae and in the granal end membranes. Unlike PS II and PS I, cyt b/ f complex is
distributed across all membrane regions (Cox and Andersson, 1981). An open question concerns a possible localization of PS I centers in the margin of the
grana stacks (Webber et al., 1988; Anderson, 1989;
Albertsson, 1995). Albertsson (2001) put forward the
hypothesis that PS I localized in the margin and ends
of the grana stacks contributes to the linear pathway
while PS I localized in the stroma lamellae contributes
to the cyclic pathway (Albertsson, 2001). The localization of PS I and PS II in different membrane regions requires long-range diffusion of the mobile carriers PQ or plastocyanin (PC). A detailed analysis of
the kinetics of electron transfer reaction between PS II
and the PQ pool has shown that diffusion of PQ is restricted to small heterogeneous domains including an
average of three to four RC, a membrane surface much
smaller than the size of a grana disk (Joliot et al., 1992;
Lavergne et al., 1992; Kirchhoff et al., 2000). It is assumed that the membrane proteins, which occupy more
than half of the membrane surface, limit the diffusion
of PQ. This implies that PQ is not involved in long distance transfer and that the linear process exclusively
involves cyt b/ f complexes localized close to PS II,
i.e., in the appressed regions. Conversely, the sole role
of the cyt b/ f complexes localized in the stroma region
might be to participate in the cyclic process (“cyclic cyt
b/ f ”).
It worth pointing out that, if the photosynthetic apparatus were exclusively devoted to a linear process,
one would expect that its optimization during evolution would have led to a random distribution of RCs
within the membrane. Such a distribution would minimize the distance between membrane proteins, leading
to faster electron exchanges mediated by PQ or PC.
Thus, the segregation of PS I and PS II centers in different membrane regions can be taken as a way to separate
the carriers involved in the cyclic and the linear flows,
which limits redox cross-talk between the cyclic and
linear chains. Structural separation between the linear
and cyclic processes is pushed to an extreme in the case
of C4 plants, in which only a cyclic process operates
in bundle sheath cells that mainly include PS I centers
(Bassi et al., 1985).
In green unicellular algae, the supramolecular organization of thylakoid membranes significantly differs from that in higher plants. Thylakoid membranes
Chapter 37
Cyclic Electron Transfer Around Photosystem I
consist of long flat vesicles (disks) that are generally
stacked in groups of 2–4, a much smaller number than
that seen in higher plants. Freeze-fracture images suggest that appressed and nonappressed regions are more
widely connected than is the case in chloroplasts of
higher plants, which are connected by narrow fret junctions. This membrane organization suggests that cyclic
and linear pathways could interact more in green algae
than in higher plants.
In cyanobacteria, thylakoids membranes are unstacked and appear as isolated flat vesicles. The organization of these membranes differs between species
but concentric arrangements of thylakoids are often seen in rod-shaped cells as Synechococcus sp.
PCC 6803 or filamentous species such as Phormidium
laminosum. There is evidence from freeze-fracture images of cyanobacterial thylakoid membranes that PS I
and PS II are typically found in the same membrane
regions — see Mullineaux (1999), although Sherman
et al. (1994) noted a slight asymmetry in the distribution of complexes in Synechococcus sp. PCC 6803,
suggesting a concentration of PS I near to the plasma
membranes. Spatial segregation of linear and cyclic
chains is thus very unlikely and one expects these pathways to share the same electron carriers. In contrast
to photosynthetic eukaryotes, thylakoids in cyanobacteria include a respiratory chain that shares the PQ
pool and the cyt b/ f complex with the photosynthetic
chain (for a review see Schmetteter, 1994). Thus, redox poising of a putative cyclic chain could be controlled by interaction with the linear photosynthetic
chain as well as with the respiratory chain. An open
question is the nature of the substrate of the NDH enzymes of the respiratory chain—NADH or NADPH—
localized in the thylakoids. NADPH-PQ reductase,
present at high concentration in the chloroplasts, could
itself contribute efficiently to a cyclic process (Fig. 1,
pathway 1).
VI. Occurrence of Cyclic Flow in
Higher Plants
Work establishing the existence of pathways for cyclic
ET has largely been performed in isolated systems, usually thylakoid membranes (broken chloroplasts), with
addition of natural or artificial mediators. While such
studies are essential in characterizing the pathway of
cyclic ET, they leave open the question; does this pathway actually operate under in vivo conditions? More
specifically, if we are to understand the function of
cyclic ET it must be established whether it occurs un-
643
der normal physiological conditions, where linear ET
is also possible.
Many of the studies that have indicated the presence
of cyclic ET in intact leaves have used rather indirect
means, typically involving conditions that largely or
totally suppress PS II turnover. While such studies are
valuable, especially in trying to understand the regulation of the cyclic pathway, they do not, in themselves,
show that this pathway is able to compete with linear
ET.
One commonly used assay taken as evidence for
cyclic flux is the measurement of the relaxation of
P700+ following illumination of a leaf with a period
of far-red light ( > 695 nm) or in the presence of a
PS II inhibitor, such as DCMU (Maxwell and Biggins,
1976). When a leaf is exposed to far-red light to oxidize P700 and then that light is abruptly cut, typically
at least two phases of P700+ reduction can be detected.
The half time for the fast phase is typically of the order
of 200–1,000 msec (Joët et al., 2002). By comparison,
in white light, under conditions where PS II is turning
over normally, the half time for P700+ reduction is of
the order of 10–20 msec. Thus, it seems immediately
unlikely that cyclic ET could ever compete effectively
with a linear flow that is 10–50 times faster. However,
measurements made in the absence of PS II turnover
will tend to result the accumulation of electron transfer components (including Fd and NADP) in the oxidized state and so, the half times measured probably
represent a gross underestimate of the maximum rate
of electron flow from the stroma to P700+ . The rate
of P700+ reduction varies between different groups of
organisms, being slow in plants and especially slow in
C3 plants (Herbert et al., 1990; Joët et al., 2002). The
rate can, however, be accelerated under certain conditions, for example under anaerobiosis (Joët et al., 2002)
or following heat stress (Maxwell and Biggins, 1976;
Burrows et al., 1998; Bukhov et al., 1999). The former
probably reflects an increase in the reduction state of the
chloroplast in the dark, increasing the supply of reductant to re-reduce P700+ . In the latter case, it is less clear
what gives rise to the effect, it may relate to a temperature induced shift in redox poise or to an “opening up”
of redox components to the surrounding medium accelerating their reduction. Recently, Golding et al. (2004),
examining the relaxation of P700+ in the presence of
the PS II inhibitor DCMU, observed that preillumination of leaves eliminates the fast component of P700+
reduction but that this effect is removed if the leaves
are experiencing drought stress. It is suggested that a
transient “cyclic-enabled” state existing in the dark is
stabilized under drought conditions (Johnson, 2005).
644
Another parameter that has been taken to indicate
cyclic ET is the presence of a transient rise in fluorescence following illumination (Asada et al., 1993;
Burrows et al., 1998). When actinic light is removed,
the fluorescence yield falls, due to the oxidation of Q−
A.
In some conditions, the yield of fluorescence is seen
to rise transiently and then fall again. This effect is attributed to the reduction of the PQ pool by electrons
originating from the stroma, indicating that a pathway
exists between the stroma and the electron transport
chain that could participate in cyclic ET. Such measurements can be made following conditions of steady-state
white light, so may reflect better the normal physiological state of the leaf but, as with the decay of P700+ ,
involve processes that are far too slow (several tens of
seconds) for them to be supposed to be involved in efficient cyclic ET in competition with linear ET. As with
the decay of P700+ following far-red light, this effect
is enhanced by exposing plants to heat stress (Sazanov
et al., 1998).
Although cyclic ET involves no net flux that can
be measured, it does nonetheless have “products” that
are measurable, notably the formation of a pH gradient across the thylakoid membrane, which can be evidenced using optical spectroscopy and the storage of
light energy, which is measured using the technique of
photoacoustics.
Two absorbance signals have been used as indicators
of the pumping of protons across the thylakoid membrane during cyclic ET: Proton translocation results
in the formation of a transmembrane electrical potential, which can be measured as an apparent absorbance
change around 515 nm; and the swelling of the thylakoid membrane that occurs when a pH is generated
induces a change in its light scattering properties, giving an apparent absorbance change in the region of
535 nm. Both of these absorbance changes can be observed when leaves are illuminated with far-red light,
indicating that cyclic ET is occurring and is generating
pH. However, the conditions needed to observe such
changes are often quite specific, so again it is difficult
to be certain whether the cycling observed is relevant
to conditions of steady-state photosynthesis.
Photoacoustics measures pressure waves produced
in samples in response to light, which can be interpreted to provide information on energy storage and
gas exchange. Photoacoustic signals are complex, with
a number of different processes contributing, so it is
necessary to design experiments carefully to give any
information on cyclic ET. As with most other methods,
this often means using far-red light to avoid any contribution from PS II photochemistry. A large number
Pierre Joliot, Anne Joliot and Giles Johnson
of studies have been published using this approach, often in combination with other approaches. For example
(Joët et al., 2002), recently combined measurements of
photoacoustics with relaxation of P700+ to investigate
the functioning of cyclic ET in tobacco.
In spite of the array of different methods that have
been applied in an attempt to determine whether cyclic
ET occurs in higher plants, the question still remains
controversial. A clear case where cyclic ET is thought
to be the norm is in the bundle sheath cells of certain
C4 plants, including maize (Herbert et al., 1990; Asada
et al., 1993; Joët et al., 2002). In such cells, PS II is
largely absent, yet these cells are responsible for the
fixation of CO2 through normal Benson–Calvin cycle
activity. By contrast, photosynthesis (and specifically
PS II) is not responsible for generating the reducing
potential required to drive CO2 fixation, since this is
generated through the oxidation and decarboxylation
of malate imported from the mesophyll. Cyclic ET is
thought to provide the ATP. Studies using photoacoustics and P700+ reduction kinetics have both provided
support for the occurrence of cyclic ET in maize bundle sheath chloroplasts (Herbert et al., 1990; Joët et al.,
2002) as have combined measurements of P700 and
chlorophyll fluorescence under combinations of far-red
and red light (Asada et al., 1993).
In C3 plants, the situation is less clear. As early as
1978, Heber et al. (1978) observed the presence of far
red-induced light scattering changes in spinach leaves.
This scattering was slow to form however and was inhibited by oxygen. The latter observation can easily be
explained in terms of the ability of oxygen to oxidize
the acceptor side of PS I. If PS II activity is suppressed,
then electrons taking part in the cyclic pathway that are
leaked to oxygen cannot be replaced. The cyclic ETC
will rapidly become completely oxidized and cyclic ET
will stop. This contrasts with the situation in C4 bundle
sheath cells, where reductant is available from the decarboxylation of malate, such that, even in the absence
of PS II activity, electrons can be reinjected in to the
cyclic pathway.
A number of studies have, however, indicated that efficient far red-induced cyclic ET can occur in C3 leaves,
however, this requires careful selection of conditions.
For example, Katona et al. (1992) observed that, under
conditions of CO2 free air, far-red light was able to efficiently energize chloroplasts in cabbage leaves, as indicated by light scattering changes. This effect was however suppressed by O2 or CO2 . Heber et al. (1992) performed similar experiments on ivy leaves and reached
similar conclusions, with the CO2 concentration again
being crucial. By contrast, the combination of red light
Chapter 37
Cyclic Electron Transfer Around Photosystem I
(rather than far-red) and low CO2 and O2 did not result
in chloroplast energization. In other words, conditions
giving rise to PS II turnover in the absence of any terminal electron acceptor result in the rapid total reduction
of the ETC. Under such conditions, no further electron
transport, linear or cyclic, is possible (see section on
redox poising above). Taking an alternative approach,
Cornic and colleagues observed the effects of periods
of actinic illumination on the ability of limiting levels
of far-red light to oxidize P700. Even short periods of
high light were found to be sufficient to reduce the effectiveness of far-red light in oxidizing P700. This was
interpreted as being due to an activation of cyclic ET
during high light, feeding electrons back into the ETC
via an efficient cyclic pathway (Cornic et al., 2000).
While the above discussion leads to the conclusion
that cyclic ET can be induced in the leaves of higher
plants, it does not tell us whether it does occur under conditions of steady-state photosynthesis. Given
the low rates of ET that have been measured and the
careful poising that is often needed to observe cyclic
ET, it is not at all clear that this pathway can compete
under conditions where linear ET is feeding electrons
into the ETC. To determine whether or not cyclic is a
real physiological phenomenon, we need to be able to
measure it under conditions of normal photosynthesis.
The most common approach evidencing cyclic ET
under conditions of steady-state photosynthesis is to
examine the relationship between PS I and PS II electron transport. Given that cyclic ET involves only PS
I and linear both PS I and PS II, any change in cyclic
ET, relative to linear ET will give a change in the ratio
of the flux of the two photosystems. Measurements of
PS II flux are usually made using analysis of chlorophyll fluorescence. The quantum efficiency of PS II is
measured as the parameter PS II (Genty et al., 1989).
Multiplying this parameter by the light intensity gives
a measure of relative flux. Provided light interception
by the PS II antenna remains constant (which might not
be the case, e.g., due to state transitions) this parameter is thought to give a robust relative measure of the
PS II electron transport rate.
In vivo measurements of PS I electron transport
under conditions of steady-state photosynthesis have
proved more controversial. A commonly used approach
has been to measure the redox state of the P700 pool
and to take the extent of reduction of this pool as a measure of the quantum efficiency of PS I. Comparisons of
PS I turnover measured in this way with PS II turnover
measured by fluorescence have been made at a variety of irradiances and CO2 concentrations (Harbinson
and Foyer, 1991; Harbinson, 1994). These studies have
645
found the relationship between these two parameters
to be linear. Thus, it has been concluded that, under
most physiological conditions, cyclic ET is either absent in the presence of light or forms a constant proportion of the linear flux. More recent data have however
found clear evidence for cyclic electron transport using
this approach (Clark and Johnson, 2001; Miyake et al.,
2004; Miyake et al., 2005a,b).
The contradictions between the above studies have
not yet been fully explained however probably relate to
the measuring conditions or the physiological status of
the plants concerned. Observations of cyclic ET have
been made under conditions where the supply of CO2 to
the leaf is restricted. For example, Harbinson and Foyer
(1991) observed that the relationship between PS I and
PS II quantum efficiency (using light as a variable) was
different in CO2 free air to that seen in the presence
of CO2 . Gerst et al. (1995) observed that, upon imposing drought stress upon a leaf in the light, PS II
was more sensitive to inhibition than PS I. By contrast,
however, in experiments where a leaf was exposed to
varying CO2 , the relationship between PS I and PS II
efficiency was found to be linear, extrapolating to the
origin (Harbinson, 1994). Thus any cyclic flow that occurs must be in proportion to linear flow. In a similar
experiment, Golding and Johnson (2003) noted that,
although the total amount of reduced P700 was proportional to PS II efficiency, this relationship did not
extrapolate to the origin, giving space for a constant
rate of cyclic ET (Golding and Johnson, 2003). In addition, however, these authors applied the method of
Klughammer and Schreiber (1994), to estimate the proportion of PS I in an “active” state. This measurement
is performed by superimposing a saturating flash of
white light on top of background actinic illumination
and then transferring the sample directly to darkness,
taking the total signal following the flash-to-dark transition as a measure of active centers. In the study of
Klughammer and Schreiber (1994), the loss of active
centers was supposed to be related to a limitation on the
acceptor side of PS I, preventing centers from turning
over. Surprisingly, Golding and Johnson (2003) noted
that active PS I rose at low CO2 (i.e., under conditions
where PS I is most likely to be acceptor-side limited).
They suggested that a distinct population of PS I centers exists that is largely or wholly involved in cyclic
ET. This “cyclic-only” pool is activated at low concentrations of CO2 and under high light. Thus, the contradictions in earlier measurements might be related
to the way in which CO2 limitation was applied and
whether these “cyclic” centers were already activated
or not.
646
In measurements of cyclic ET in far-red light it has
been common to take the rate of P700+ reduction as
an indicator of PS I turnover (Maxwell and Biggins,
1976). Essentially, the same approach can be applied
in white light conditions. This relies on the limiting step
in electron transport lying prior to PS I, which is usually the case under physiological conditions, such that
the flow of electrons to P700 can be measured and reflects the overall flux through PS I. A potential problem
with this method was noted by Sacksteder and Kramer
(2000), who noted that a net reduction of cyt f , measured at the time the light is switched off, might have
to be taken into account to determine the electron flux
toward P700 (Sacksteder and Kramer, 2000). Strictly,
there should be no net reduction of cyt f at the moment
when the light is switched off, as at steady-state the rate
of oxidation and reduction are identical and neither of
these are instantly affected by cutting the light. Practically, given the time resolution and sensitivity of most
instruments, this may be a problem but only under low
light conditions where P700 is largely reduced but cyt
f oxidized, giving rise to a short time lag in the reduction of cyt f . Sacksteder and Kramer (2000) have
compared the turnover of PS I and PS II in greenhouse
grown sunflowers. They observed a linear relationship
between PS I and PS II ET, implying no (or a constant proportion of) cyclic ET, the same conclusion as
reached by Harbinson and colleagues using P700 redox state to measure PS I turnover (Harbinson et al.,
1990; Harbinson, 1994). In contrast, Clarke and Johnson compared the rates of PS I and PS II turnover across
a range of temperatures and light intensities in barley
grown in a growth cabinet. They observed that PS II
photochemistry saturated at lower irradiances and was
more sensitive to low temperature than PS I. Thus, they
concluded that high light and low temperatures lead to
enhancement of cyclic ET (Clarke and Johnson, 2001).
The contradiction between these two studies may reflect a species difference, but is more likely explained
by the range of light intensities used in each case. In
the experiments of Sacksteder and Kramer, the highest light intensities used were just saturating, whereas
for Clarke and Johnson light intensities were used that
went well above saturating for PS II electron transport
(though not necessarily for PS I). Thus it appears that
cyclic ET is a characteristic of saturating light, although
a low rate at subsaturating light cannot be excluded, if
this forms a constant proportion of the linear flux. A
similar approach by Golding and Johnson (2003) drew
the same conclusion concerning responses to low CO2
and drought.
A new technical approach, based on membrane potential measurements (Joliot and Joliot, 2002, 2004)
Pierre Joliot, Anne Joliot and Giles Johnson
has been developed to determine the absolute rate
of the cyclic and linear pathways whatever the intensity of illumination. In this method, the sum of
the rates of photochemical reactions I and II is measured by the difference in the rate of membrane potential changes determined immediately before or after
switching off the light. Experiments were performed
under strong light excitation in the presence of air, with
dark-adapted spinach (Joliot and Joliot, 2002) or Arabidopsis thaliana leaves (Joliot and Joliot, 2004), i.e.,
in conditions where the Benson–Calvin cycle and thus,
the linear ET, is mainly inactive. Under saturating illumination, rate of the cyclic flow is estimated to ∼130
sec−1 and remains roughly constant during the first 10
sec of illumination. Unexpectedly, this cyclic process
is not inhibited by antimycin (Joliot and Joliot, 2002).
Under the same conditions, fluorescence induction kinetics shows that a pool of soluble PS I acceptors (Fd,
FNR and NADP) of approximately nine electron equivalents is reduced in less than 100 msec via the linear
pathway. This implies that, even in dark-adapted leaves,
PS I remains able to transfer electrons to a pool of oxidized PS I acceptors. In the presence of DCMU, where
the linear flow is fully inhibited, a similar rate of the
cyclic flow is measured during the first seconds of illumination. This rate progressively drops to zero in ∼7
sec, due to slow electron leaks that lead to the oxidation
of the carriers involved in the cyclic chain.
After 100 msec of illumination at an intensity that
is saturating for the cyclic process, most of P700 is
reduced (Harbinson and Hedley, 1993; Strasser et al.,
2001; Joliot and Joliot, 2002; Schankser et al., 2003)
implying that a fast charge recombination between
P700+ and reduced acceptors will occur. In agreement
with this assumption, it is observed that the kinetics of
the membrane potential displays a fast decaying phase
of small amplitude, which is completed in ∼ 500 μsec.
The amplitude of this phase is roughly proportional
to the light intensity. This phase correlates with a reduction phase of P700+ (measured by the absorption
changes at 810 nm) and has been ascribed to a charge
recombination between P700+ and most probably the
iron-sulfur carrier FX (Joliot and Joliot, 2002). It can
be thus concluded that most of the carriers of the linear
and cyclic chains are poised in their reduced state. A
small fraction of “cyclic PS I” centers includes P700+
and a single negative charge on the FA /FB cluster. For
these RCs, the rate of P700+ reduction via the cyclic
pathway is faster than the rate of charge recombination
between P700+ and (FA /FB )− (t1/2 ∼ 45 msec; Hiyama
and Ke, 1971).
A charge recombination process is also observed in
the presence of DCMU, the amplitude of which is half
Chapter 37
Cyclic Electron Transfer Around Photosystem I
that measured in its absence. This is explained by the
fact that illumination induces the oxidation of all the
carriers of the linear chain, while the carriers involved
in the cyclic pathway are transitorily poised at a potential able to induce the reduction of most of the FA /FB
acceptors (<−600 mV). In the 1–7 sec time range, the
decrease in the rate of the cyclic flow is associated with
a corresponding decrease in the amplitude of the charge
recombination phase. This parallel decrease reveals a
progressive oxidation of all the carriers involved in the
cyclic process, associated with a slow electron leak toward O2 or the Benson–Calvin cycle.
VII. Pathway of Cyclic Flow in Higher
Plants
Work described above leads us to conclude that cyclic
ET is a real physiological phenomenon, though perhaps only occurring under a limited range of conditions.
The question then arises, what is (are) the physiological pathway(s) for this electron flow. Early work indicating two pathways of cyclic ET in isolated systems
were mainly based on measurements of chloroplasts
isolated from C3 plants. We would therefore expect to
see evidence for both the ferredoxin-linked antimycinsensitive and the NADPH-linked antimycin-insensitive
pathways in whole leaves. The in vivo data on the
effect of antimycin A are however, ambiguous. Joët
et al. (2002) reported a slow cyclic ET measured under
far-red excitation is stimulated in anaerobiosis but also
in the presence of inhibitors of the respiratory chain,
including antimycin. In the same way, antimycin does
not inhibit the fast cyclic flow measured under strong
illumination of dark-adapted leaves (Joliot and Joliot,
2002). In both cases, it was proposed that the effect
of antimycin is related to the inhibition of the respiratory chain, which increases the reducing power in
chloroplasts via mitochondria–chloroplast interactions. Further confusion might arise in in vivo measurements because of other functions of antimycin, notably its ability to inhibit nonphotochemical quenching
(Oxborough and Horton, 1987). We thus conclude that
effect of antimycin is not a decisive test in proposing
or excluding the occurrence of cyclic electron flow.
A. NADPH-Dependent Cyclic Electron
Transfer
The more convincing arguments that favor the involvement of NADPH:PQ reductase (NDH) in the cyclic
pathway come from the analysis of mutants lacking
this enzyme. NDH-dependent cyclic ET reported in the
647
literature appears as a slow process that is very likely related to the low concentration of NDH [∼1% of that of
the photosynthetic chain (Sazanov et al., 1995)]. Levels
of this complex are seen to increase under conditions
of stress, suggesting a role in survival under conditions
of oxidative stress (Casano et al., 2000; Teicher et al.,
2000; Lascano et al., 2003), however, to what extent this
increases the flux through the NDH complex remains
unclear. Burrows et al. (1998) observed that plants
lacking the NDH complex lacked the transient fluorescence rise seen following illumination. Hashimoto
et al. (2003) were recently able to use this characteristic to identify a new mutant of A. thaliana, CRR2,
which lacks a nuclear encoded factor that seems to be
essential for NDH expression. However, the absence
of a fluorescence rise in NDH-less mutants seems to
be highly sensitive to the developmental and metabolic
status of the leaf (A. Krieger-Liszkay, personal communication.), supporting the idea that this is not the
only route for reduction of the PQ pool by stromal
factors. Evidence has also been published that the reduction of P700+ following far-red light is affected in
plants lacking the NDH complex (Joët et al., 2002).
Involvement of NDH in the fast cyclic ET measured in
dark-adapted leaves submitted to strong illumination
(Joliot and Joliot, 2002, 2004) appears unlikely as this
enzyme, which is present at low concentration, would
have to operate at rate as high as ∼104 sec−1 to sustain
a rate of electron flow of 130 sec−1 .
B. Ferredoxin-Dependent Cyclic Electron
Transfer
If rapid rates of cyclic ET cannot be sustained by the
NDH complex, then we must consider the likely involvement of a ferredoxin pathway. Electrons maybe
directly transferred from Fd either directly to cyt b/ f
(Fig. 1, pathway 3) or via a FQR (Fig. 1, pathway 2;
Bendall and Manasse, 1995). In Fig. 2, a mechanism
is proposed that results in the pumping of one proton
per electron transferred, but differs from conventional
Q-cycle process (Joliot and Joliot, 2004). FNR is
known to copurify with the cyt b/ f complex (Clark
et al., 1984) and provides a binding site for Fd (Zhang
et al., 2001). It seems likely that the electron pathway between Fd and site Qi involves the covalently
bound cyt c (cyt ci ) recently characterized in the structure of cyt b/ f complex in Chlamydomonas reinhardtii
(Stroebel et al., 2003) and in cyanobacteria (Kurisu
et al., 2003). On the other hand, it has been proposed
that an electron carrier G (Lavergne, 1983; Joliot and
Joliot, 1988), is localized on the stromal side of the cyt
b/ f complex and able to exchange electrons with cyt
648
Fig. 2. A mechanism for the ferredoxin-dependent electron
transfer.
bh . This carrier G has been now identified as cyt ci
(Alric et al., 2005). According to Fig. 2, sites Qi and
Qo behave quite symmetrically. At site Qi , reduction
of PQ involves the transfer of one electron from Fd
and one electron from cyt bh ; at site Qo , oxidation of
PQH2 involves the transfer of one electron to cyt f via
the Rieske Fe/S protein and one electron to cyt bl . It
worth noting that, in the dark, reduction of the PQ pool
could be induced by the sequential transfer of electrons
from Fd to site Qi and the cyt b/ f complex would then
behave as a FQR. Evidence for a distinct FQR, independent of the Qi site, is related to the effect of antimycin
compared to HQNO (Moss and Bendall, 1984). This
evidence only rules out a role for antimycin in blocking
the Qi site and does not rule out the involvement of that
site, if it does not bind antimycin.
It has been proposed that the small hydrophilic
polypeptide PGR5 is required in a process of proton
gradient generation and could be involved in FQR activity (Munekage et al., 2002). At present, very little
information on this mutant is available and a better
Pierre Joliot, Anne Joliot and Giles Johnson
characterization of its function is required. PGR5 lacks
a metal binding motif that might implicate it in a direct
role in electron transfer, however it might play a secondary role, e.g., in being involved in the binding of
FNR to the cyt b/ f complex. The published evidence
that PGR5 is involved in cyclic ET is based on the observation that NADPH and Fd-dependent reduction of PQ
pool, measured by chlorophyll fluorescence is impaired
in PGR5 mutant. This observation needs to be considered with care, however, since the effect on the kinetics
of PQ reduction and the sensitivity to antimycin appear
to be qualitatively the same, with only the amplitude of
the fluorescence rise being changed (Munekage et al.,
2002). Since the latter might be sensitive to other factors, for example the presence of photoinhibited PS II
centers in the thylakoid membrane, this result is not
conclusive.
The fraction of PS I centers that operate according
to the cyclic or linear mode has been determined by
measuring P700 and PC oxidation under weak far-red
excitation (8 photons / PSI / s), a photochemical rate
constant well below the rate constant of the limiting
steps of cyclic or linear flow (Joliot and Joliot, 2005).
Far-red illumination of a dark-adapted leaf induces a
slow oxidation of P700 and PC that is completed in
10–20 sec. During this oxidation phase, the number of
PS I turnovers is much larger than the number of electrons stored in the primary and secondary PS I donors.
It implies that most of the electrons formed on the stromal side of PS I are transferred back to PS I via the
cyt b/ f complex (cyclic electron flow). Slow electron
leaks, probably toward the Benson-Calvin cycle, result
in a progressive oxidation of most of the carriers involved in the cyclic pathway, including P700. When the
same leaf is first preilluminated for several minutes under green light that excites both photosystems and activates the Benson-Calvin cycle, P700 oxidation induced
by far-red excitation is a much faster process completed
in less than 3 sec. The kinetics of P700 oxidation measured with a preilluminated leaf is close to that measured in the presence of methyl viologen, an efficient
PS I electron acceptor. It implies that, in preilluminated leaves, most of the electrons reaching the stromal
size of PS I are transferred to the Benson-Calvin cycle via FNR and NADP (linear electron flow). Using
the same approach, the transition from a cyclic (darkadapted leaf) to a linear mode (preilluminated leaf) has
been recently analyzed during the course of the green
preillumination (P. Joliot, unpublished data). For the
first minute of illumination, most of PS I centers contribute to cyclic electron flow. During the subsequent
period of illumination (1–10 min), activation of the
Chapter 37
Cyclic Electron Transfer Around Photosystem I
649
Fig. 3. A model for the structural organization of linear and cyclic ways within the photosynthetic membrane.
Benson-Calvin cycle, as shown by the increase in the
rate of linear flow, correlates with a decrease in the rate
of the cyclic flow. In the same way, the slowdown of
the linear flow induced by a lack of CO2 is associated
with an increase of the rate of the cyclic flow, as previously proposed by Heber and co-workers (Hauser et al.,
1995). These results suggest that linear and cyclic pathways be in permanent competition for the reoxidation
of Fd. The relative efficiency of cyclic and linear pathways is suggested to be determined by the probability of
Fd binding either to a site localized on the stromal site
of the cyt b/ f complex or to an ‘active’ FNR. This we
define as an FNR molecule to which NADP+ is bound.
Activation of cyclic flow is thus associated with the reduction of NADP+ and the competition between cyclic
and linear mode will be controlled by the redox state
of NADP which depends upon the degree of activation
of the Benson-Calvin cycle. In dark-adapted leaves, illumination first induces the reduction of the small pool
of acceptors present in their oxidized state (FNR and
NADP+ ). When this pool is fully reduced, Fd in excess
is available to bind the stromal site of cyt b/ f , which
initiates cyclic electron flow.
In the model depicted in Fig. 3, Fd is presumed to
freely diffuse in the stromal compartment. This contrasts with models in which the cyclic chain is organized in supercomplexes that associate 1 PS I center / a
cyt b/ f 1 PC / 1 Fd (Carillo and Vallejos, 1983; Laisk
et al., 1992; Laisk, 1993; Joliot and Joliot, 2002). In
this hypothesis, the concentration of PS I included in
the supercomplex cannot be larger than that of cyt b/ f
complex localized in the non-appressed region of the
membrane. As most of PS I can be involved in cyclic
electron flow (Joliot and Joliot, 2005) and only ∼50%
of cyt b/ f complex is localized in the non-appressed region Albertsson, 2001), the obligatory formation of supercomplexes for cyclic electron flow can be excluded.
VIII. Cyclic Flow in Green Unicellular
Algae
There is a general consensus that a cyclic electron flow
operates in algae, at least under anaerobic conditions,
when PS II activity is mainly inhibited due to the reduction of QA . It worth pointing out that, in contrast to
higher plants, anaerobiosis is a common occurrence for
many algae in their natural environment. Anaerobiosis
is known to induce state 1 to state 2 transition associated with changes in the distribution of the membrane
complexes (Bulté et al., 1990) to a much larger extent
than those induced by chromatic adaptation (Bonaventura and Myers, 1969). Anaerobiosis, like all treatments
that decrease the ATP content of the cell (mitochondrial inhibitors, uncouplers, ATP-synthase inhibitors),
induces the transfer of most of the light harvesting complex LHC II (Bulté et al., 1990) and of a fraction of
cyt b/ f complexes (Vallon et al., 1991) from the appressed to the nonappressed region of the thylakoids.
On the basis of measurements of the turnover rate of
cyt f , Finazzi et al. (1999) concluded that only linear
ET is active in the presence of O2 (state 1 conditions),
while cyclic electron flow operates in state 2 conditions (anaerobiosis or presence of uncouplers), where
the linear process is fully inhibited. The rate of cyclic
flow measured in state 2 conditions is similar to that
650
measured for the linear flow at the same light intensity in state 1 conditions. One can thus conclude that in
anaerobic conditions, most of the antennae is involved
in the cyclic process with ∼80% of the light collected
by the “cyclic PS I.” In these conditions, unicellular
algae may behave in a similar way to the green bacteria with type I photosystems, with a photosynthetic
process entirely devoted to ATP synthesis. The cyclic
process in algae thus appears fundamentally different
from that in higher plants, which occurs in the presence
of O2 and state 1 conditions.
A. Mechanisms of Cyclic Flow in Algae
Complexes containing PS I and cyt b/ f complexes
have been observed in solubilized membranes of
C. reinhardtii (Wollman and Bulté, 1989) possibly
pointing to the presence of supercomplexes in this organism. On the other hand, the analysis of ET kinetics
in state 2 conditions in a mutant with a low cyt b/ f content has shown that PC is able to freely diffuse from any
PS I center to cyt f (Finazzi et al., 2002). This excludes
the presence of functional supercomplexes, including
a trapped PC. In state 2 conditions, a spatial separation between the linear and cyclic chains results from
the transfer of a large fraction of the stromal cyt b/ f
complexes from the appressed to the nonappressed region. Thus, one expects that, even in the absence of
supercomplexes, an efficient cyclic ET will occur in
the nonappressed region that includes the PS I centers
and most of the cyt b/ f complexes.
The electron pathway from PS I to the cyt b/ f complex has not been identified. While no gene encoding
for an enzyme similar to NDH has been identified in
the chloroplast of algae, the presence of a PQ reductase is suggested by the slow reduction of PQ pool
in the appressed region, which has been observed in
the presence of O2 (Diner and Mauzerall, 1973). One
expects that such an enzyme would operate at a much
higher rate in anaerobic conditions. This enzyme could
be involved in a cyclic process if it were present in the
nonappressed region. Another possible pathway is a direct ET from PS I to cyt b/ f complex, as depicted in
Fig. 1 (pathway 3). It worth pointing out that, to our
knowledge, complexes that associate FNR and the cyt
b/ f complex have not been identified in C. reinhardtii.
IX. Cyclic Flow in Cyanobacteria
In cyanobacteria, the presence of respiratory and photosynthetic electron transport chains in the same mem-
Pierre Joliot, Anne Joliot and Giles Johnson
brane system, to the point where the two share redox
components (Scherer, 1990), means that the cyclic ET
occurs in an environment very different to that in plants
and algae. A common feature to most of cyanobacteriae is the large excess of PS I as compared to PS II
centers (Fujita et al., 1994). This suggests that even
under steady-state condition of illumination, a fraction of PS I centers could be involved in cyclic ET.
On the other hand, in cyanobacteria, PS I centers are
organized in trimers, which exclude the formation of
supercomplexes that associate PS I centers with the cyt
b/ f complex.
After illumination in the presence of DCMU or after far-red excitation, reduction of P700+ occurs with
t1/2 ∼ 500 msec, taken to be indicative of cyclic ET
(Maxwell and Biggins, 1976; Mi et al., 1992a,b; Yu et
al., 1993). At the same time, measurements of energy
storage via photoacoustics point to the occurrence of
cyclic ET under such conditions. However, it can be assumed that, in dark-adapted samples, the PQ pool will
be maintained in a relatively reduced state due to respiratory electron flow. It needs to be shown, therefore, that
electron transport from the acceptor side of PS I back
into the PQ pool occurs and is maintained under conditions where PS II is active. (Sandmann and Malkin,
1983, 1984) were able to observe that, while respiration was inhibited in the light, due to competition with
photosynthesis, NAD(P)H oxidation by spheroplasts
from Aphanocapsa was less affected. The addition of
DCMU did not enhance NAD(P)H oxidation, suggesting no competition between NDH and PS II for PQ
reduction. However, as pointed out by Mi et al. (2000),
the organization of the membrane systems in cyanobacteria may be substantially disturbed in isolated systems.
As far as we are aware, these results have not yet been
confirmed in intact cells.
Measurements on mutants lacking the NDH complex suggest this is generally the primary route for
cyclic ET (Yu et al., 1993; Mi et al., 2000). Mutants
lacking ndhF of the NDH complex and psaE in PS
I, show inhibited growth at low-light intensities, but
are unaffected at high light. Cyclic ET in cyanobacteria has been implicated in providing energy for CO2
concentrating mechanisms. Growth of Synechocystis
at low CO2 results in upregulation of NDH subunits
(Deng et al., 2003). Mutants lacking the NDH complex have a reduced ability to tolerate growth under
limiting CO2 conditions (Ogawa, 1991). On the other
hand, it has been reported that Synechocystis sp. PCC
6803 contains a succinate quinol oxidoreductase that is
suggested to contribute to cyclic electron flow (Cooley
et al., 2000). This enzymatic activity would, however,
Chapter 37
Cyclic Electron Transfer Around Photosystem I
involve a more complex cyclic pathway than so far
considered and it is questionable whether it can still be
regarded as cyclic electron flow.
Exposure to high salt concentrations leads to an activation of a cyclic ET pathway that does not require
NDH (Jeanjean et al., 1998). The details of this pathway
remain unclear but probably involve FNR (van Thor
et al., 2000; Matthijs et al., 2002). Binding of the FNR
to the membrane is facilitated by an N-terminal domain
which is absent in the higher plant enzyme (van Thor
et al., 2000). Low concentrations of sodium bisulfite
have also recently been suggested to stimulate cyclic
ET (Wang et al., 2003).
X. Functions and Regulation of Cyclic
Electron Transfer
The primary role of cyclic ET has always been supposed
to be the generation of ATP, either to support CO2 fixation or for other metabolic processes. The requirement
for cyclic ET to balance the production of reductant
and ATP in normal CO2 fixation conditions has been
widely debated. Structural analysis of the electrical rotor of the chloroplast ATP-synthase has shown that it is
made up of 14 polypeptides (Seelert et al., 2000). One
thus expects that a complete rotation of the rotor, which
induces the synthesis of 3 ATP, is associated with the
transfer of 14 protons across the membrane that leads
to H+ /ATP ratio of 14/3 = 4.7. Assuming that 3 protons
are pumped per electron transferred via the linear chain,
the number of ATP synthesized per electron is 3/4.7 =
0.64 or ∼2.55 ATP per CO2 , a value over-estimated as
ion leaks through the membrane are not taken into account. This amount of ATP is definitely lower than that
required by the Benson–Calvin cycle (3 ATP/CO2 ). It
is thus likely that, even in light-adapted conditions, a
small fraction of PS I contributes to a cyclic flow to
satisfy the ATP requirement of the Benson–Calvin cycle. This could explain why the minimum quantum requirement of the photosynthetic process measured under weak excitation (10–12 quanta/O2 ) is significantly
larger than the theoretical value of eight quanta/O2 .
If cyclic ET is contributing ATP to drive CO2 fixation
and assuming that this is regulated in a way that ensures
that the ATP supply is maintained at a constant level,
then we would expect that under steady-state condition
of illumination, the rate of cyclic ET would parallel the
rate of linear ET. Thus, it is perhaps unsurprising that
light-limited steady-state measurements, which rely on
comparing PS I and PS II ET, have typically found no
evidence for this cyclic flow (Harbinson et al., 1990;
651
Sacksteder and Kramer, 2000). By contrast, when the
ATP consumption is not directly coupled to CO2 fixation, we would expect to find discrepancies between
PS I and PS II turnover. This would certainly be the
case in C4 bundle sheath cells, where the reductant required for CO2 fixation is not provided by linear ET. It
would also be the case during the first few seconds of
illumination, during which time a trans-thylakoid pH
gradient must be established before ATP synthesis can
begin (Joliot and Joliot, 2002).
In green algae, reducing conditions (anaerobiosis)
result in a large state transition, with phosphorylation
of LHC II resulting in its migration to PS I, pushing the chloroplast into a “cyclic-only” state. Under
anaerobiosis, mitochondrial respiration will be inhibited, leading to a deficit of ATP and possibly also to a
rise in the concentration of NADH. A shift from linear
to cyclic photosynthetic ET would counteract this effect, lowering the production of reducing potential in
favor of ATP synthesis. Thus the redox regulation of
state transitions in algae and the resultant regulation
of cyclic ET is a mechanism for balancing the overall
cellular ATP/NAD(P)H ratio.
In higher plants, state transitions are less prominent
and their regulation in response to redox potential very
different. Reducing conditions induced by anaerobiosis do not give rise to state 1 to state 2 transitions but
they are inhibited under conditions, where the Benson–
Calvin cycle is active, though the action of thioredoxin
(Aro and Ohad, 2003). On the other hand, and in contrast to algae, efficient cyclic ET operates during the
first seconds of illumination of dark-adapted leaves (Joliot and Joliot, 2002), i.e., in pure state 1 conditions,
and there is no evidence that state transitions lead to
an increase in cyclic ET, rather they are thought to balance excitation of PS I and PS II for optimal linear
ET. The small state transitions that do occur are seen
at very low light and depend on the redox state of the
PQ pool. On the other hand, the observation that A.
thaliana grown under very low-light increase PS I relative to PS II might suggest a role for cyclic ET under
such conditions, implying a switch from CO2 fixation
to ATP synthesis (Bailey et al., 2001).
A notable difference between higher plants, some
algae and cyanobacteria is in their ability to protect
themselves from high light through the process of high
energy state quenching (qE). This process, which is
linked to the presence of the carotenoid zeaxanthin,
is driven by pH. It was first suggested some years
ago that the pH required to generate this quenching
was generated by cyclic ET (Heber and Walker, 1992);
however, it is only recently that direct evidence for its
652
requirement has been obtained. The PGR5 mutant of A.
thaliana that is thought to be deficient in cyclic ET, was
isolated using a screen that selected for a deficiency
in quenching (Munekage et al., 2002). Golding and
Johnson observed a quantitative link between the extent
of non-photochemical quenching (NPQ) and the extent
of cyclic ET under conditions of low CO2 and drought
(Golding and Johnson, 2003).
Given the dual function of cyclic ET in higher plants,
it is tempting to suggest that the different pathways
of cyclic ET may be fulfilling different functions and
therefore be differentially regulated. Upregulation of
cyclic at low CO2 correlates with a downregulation of
the cyt b/ f complex, inhibiting linear ET (Golding and
Johnson, 2003). This downregulation of linear ET has
been suggested to be regulated via thioredoxin (Johnson, 2003). Thus, upregulation of cyclic ET in response
to reducing conditions seems likely, although the mechanism involved is different to that seen in green algae.
Joliot and Joliot (2002) suggested that the cyclic ET
seen following a dark to light transition in dark-adapted
leaves maybe regulated by the ATP/ADP ratio in the
chloroplast. This is a reasonable suggestion, if we accept that this cyclic is required to kick-start ATP synthesis. On the other hand, the cyclic ET seen under such
conditions has a short lag period, during which time the
Fd and NADPH pools are likely to become reduced,
in agreement with a redox regulation process. After a
preillumination, the transition from the linear to the
cyclic mode requires more than 1 hour of dark adaptation (Golding et al., 2004; Joliot and Joliot, 2005).
Deactivation of the Benson-Calvin cyclic following illumination is slow, taking up to 3 hours or more in some
species. Thus the long period of time required for the
induction of a cyclic activated state might be explained
simply by this deactivation, though changes in the energy status of the cell, related to the consumption of
carbohydrate reserves might also play a role.
Given the low requirement for additional pH arising from CO2 fixation, it might be speculated that the
capacity of NDH pathway is sufficient to support this.
However, mutants lacking NDH are clearly not deficient in CO2 fixation. It may be however that a normally stress (redox) activated cyclic pathway becomes
partially activated under conditions where NDH is lacking, compensating for the lack of this pathway. There
is little evidence that the NDH pathway contributes to
photoprotection. Ndh-less mutants have normal levels of NPQ and are not more susceptible to photoinhibition either of PS I or PS II at ambient or low
temperature (Barth and Krause, 2002). On the other
hand, there are some indications that plants lacking
Pierre Joliot, Anne Joliot and Giles Johnson
NDH are more sensitive to water stress (Horvath et al.,
2000).
In cyanobacteria, there is a large amount of data that
link the NDH complex to CO2 concentrating mechanisms. It seems that there are distinct forms of the
complex that are involved in this process, with there
possibly being a direct role of NDH in the formation of
HCO−
3 from CO2 . It is not clear how this relates cyclic
ET however, i.e., whether this reaction depends on electron transport occurring through PS I (see discussion
in Badger and Price, 2003).
XI. Conclusion
The pathway of linear electron transport has been
clearly established for many years. Cyclic ET on the
other hand remains enigmatic. Nevertheless, this is an
area in which significant progress is beginning to be
made. The establishment of new methods for the quantitation of cyclic flow, such as those outlined here, will
help in this progress. Clearer ideas on the function of
cyclic ET will also help in the identification of the proteins involved in this pathway, either through conventional screening approaches or through reverse genetics, examining the effects of knockout mutations on
cyclic electron transport.
Acknowledgments
We would like to thank Dr. Giovanni Finazzi (IBPC,
Paris, France) and Dr. Conrad Mullineaux (University College, London, UK) for useful discussions and
Prof. Toshiharu Shikanai (Nara Institute of Science and
Technology, Japan) for providing a reprint of his work.
References
Albertsson PA (1995) The structure and function of the chloroplast photosynthetic membrane – a model for the domain organization. Photosynth Res 46: 141–149
Albertsson PA (2001) A quantitative model of the domain structure of the photosynthetic membrane. Trends Plant Sci 6: 349–
354
Alric J, Pierre Y, Picot D, Lavergne J and Rappaport F (2005)
Spectral and redox characterization of the heme ci of the cytochrome b6 f complex. Proc Natl Acad Sci USA 102: 15860–
15865
Allen JF (1983) Regulation of photosynthetic phosphorylation.
CRC Crit Rev Plant Sci 1: 1–22
Allen JF (2003) Cyclic, pseudocyclic and noncyclic photophosphorylation: new links in the chain. Trends Plant Sci 8: 15–19
Chapter 37
Cyclic Electron Transfer Around Photosystem I
Anderson JM (1989) The grana margins of plant thylakoid membranes. Physiol Plant 76: 243–248
Andersson B and Anderson JM (1980) Lateral heterogeneity in
the distribution of chlorophyll–protein complexes of the thylakoid membranes of spinach chloroplasts. Biochim Biophys
Acta 593: 427–440
Arnon DI (1955) The chloroplast as a complete photosynthetic
unit. Science 122: 9–16
Arnon DI (1959) Conversion of light into chemical energy in
photosynthesis. Nature 184: 10–21
Arnon DI and Chain RK (1977) Role of oxygen in ferredoxincatalyzed cyclic photophosphorylation. FEBS Lett 82: 297–
302
Arnon DI, Allen MB and Whatley FR (1954) Photosynthesis by
isolated chloroplasts. Nature 174: 394–396
Arnon DI, Losada M, Nozaki M and Tagawa K (1961) Photoproduction of hydrogen, photofixation of nitrogen and a unified
concept of photosynthesis. Nature 190: 601–606
Aro EM and Ohad I (2003) Redox regulation of thylakoid protein
phosphorylation. Antioxid Redox Signal 5: 55–67
Asada K, Heber U and Schreiber U (1993) Electron flow to the
intersystem chain from stromal components and cyclic electron flow in maize chloroplasts, as detected in intact leaves
by monitoring redox change of P700 and chlorophyll fluorescence. Plant Cell Physiol 34: 39–50
Badger M and Price GD (2003) CO2 concentrating mechanisms
in cyanobacteria: molecular components, their diversity and
evolution. J Exp Botany 54: 609–622
Bailey S, Walters RG, Jansson S and Horton P (2001) Acclimation of Arabidopsis thaliana to the light environment: the
existence of separate low light and high light responses. Planta
213: 794–801
Barth C and Krause GH (2002) Study of tobacco transformants
to assess the role of chloroplastic NAD(P)H dehydrogenase in
photoprotection of photosystems I and II. Planta 216: 273–279
Bassi R, dal Belin Peruffo A, Barbato R and Ghisi R (1985) Differences in chlorophyll–protein complexes and composition
of polypeptides between thylakoids from bundle sheaths and
mesophyll cells in maize. Eur J Biochem 146: 589–595
Bendall DS and Manasse RS (1995) Cyclic photophosphorylation and electron transport. Biochim Biophys Acta 1229:
23–38
Bonaventura C and Myers J (1969) Fluorescence and oxygen
evolution from Chlorella pyrenoidosa. Biochim Biophys Acta
189: 366–383
Bukhov NG, Wiese C, Neimanis S and Heber U (1999) Heat
sensitivity of chloroplasts and leaves: leakage of protons from
thylakoids and reversible activation of cyclic electron transport. Photosynth Res 59: 81–93
Bulté L, Gans P, Rebéillé F and Wollman F-A (1990) ATP control
on state transitions in vivo in Chlamydomonas reinhardtii.
Biochim Biophys Acta 1020: 72–80
Burrows PA, Sazanov LA, Svab Z, Maliga P and Nixon PJ
(1998) Identification of a functional respiratory complex in
chloroplasts through analysis of tobacco mutants containing
disrupted plastid ndh genes. EMBO J 17: 868–876
Carillo N and Vallejos RH (1983) The light-dependent modulation of photosynthetic electron transport. TIBS February 1983:
52–56
Casano LM, Zapata JM, Martin M and Sabater B (2000)
Chlororespiration and poising of cyclic electron transport –
plastoquinone as electron transporter between thylakoid
653
NADH dehydrogenase and peroxidase. J Biol Chem 275: 942–
948
Clark RD, Hawkesford MJ, Coughlan SJ, Bennett J and Hind
G (1984) Association of ferredoxin NADP+ oxidoreductase
with the chloroplast cytochrome b–f complex. FEBS Lett 174:
137–142
Clarke JE and Johnson GN (2001) In vivo temperature dependence of cyclic and pseudocyclic electron transport in barley.
Planta 212: 808–816
Cleland RE and Bendall DS (1992) Photosystem-I cyclic electron
transport – measurement of ferredoxin-plastoquinone reductase activity. Photosynth Res 34: 409–418
Cooley JW, Howitt CA and Vermaas WFJ (2000) Succinate:quinol oxidoreductase in the cyanobacterium Synechocystis sp. Strain PCC 6803: presence and function in
metabolism and electron transport. J Bacteriol 182: 714–722
Cornic G, Bukhov NG, Wiese C, Bligny R and Heber U (2000)
Flexible coupling between light-dependent electron and vectorial proton transport in illuminated leaves of C-3 plants. Role
of photosystem I-dependent proton pumping. Planta 210: 468–
477
Cox RP and Andersson B (1981) Lateral and transverse organisation of cytochromes in the chloroplast thylakoid membrane.
Biochem Biophys Res Commun 103: 1336–1342
Crofts AR, Meinahrdt SW, Jones KR and Snozzi M (1983) The
role of the quinone pool in the cyclic electron-transfer chain of
Rhodopseudomonas sphaeroides. A modified Q-cycle mechanism. Biochim Biophys Acta 723: 202–218
Deng Y, Ye JY and Mi H (2003) Effects of low CO2 on NAD(P)H
dehydrogenase, a mediator of cyclic electron transport around
Photosystem I in the cyanobacterium Synechocystis PCC
6803. Plant Cell Physiol 44: 534–540
Diner B and Mauzerall D (1973) Feedback controlling oxygen
production in a cross-reaction between two photosystems in
photosynthesis. Biochim Biophys Acta 305: 329–352
Finazzi G, Furia A, Barbagallo RP and Forti G (1999) State transitions, cyclic and linear electron transport and photophosphorylation in Chlamydomonas reinhardtii. Biochim Biophys
Acta 1413: 117–129
Finazzi G, Rappaport F, Furia A, Fleischmann M, Rochaix JD,
Zito F and Forti G (2002) Involvement of state transitions in
the switch between linear and cyclic electron flow in Chlamydomonas reinhardtii. EMBO Rep 3: 280–285
Fork DC and Herbert SK (1993) Electron transport and photophosphorylation by Photosystem I in vivo in plants and
cyanobacteria. Photosynth Res 36: 149–168
Frenkel AW (1954) Light induced phosphorylation by cell-free
preparations of photosynthetic bacteria. J Am Chem Soc 76:
5568–5569
Fujita Y, Murakami A, Aizawa K and Ohki K (1994) Shortterm and long-term adaptation of the photosynthetic apparatus: homeostatic properties of thylakoids. In: Bryant DA (ed)
The Molecular Biology of Cyanobacteriae, Vol 1, pp 677–692.
Kluwer Academic Publishers, Dordrecht
Genty B, Briantais J-M and Baker NR (1989) The relationship between the quantum yield of photosynthetic electron transport
and quenching of chlorophyll fluorescence. Biochim Biophys
Acta 990: 87–92
Gerst U, Schreiber U, Neimanis S and Heber U (1995) Photosystem I dependent cyclic electron flow contributes to control
of Photosystem II in leaves when stomata close under water stress. In: Mathis P (ed) Photosynthesis: From Light to
654
Biosphere, Proceedings of the Xth International Photosynthesis Congress, Vol II, pp 835–838. Kluwer Academic Publishers, Montpellier
Golding AJ and Johnson GN (2003) Down-regulation of linear and activation of cyclic electron transport during drought.
Planta 218: 107–114
Golding AJ, Finazzi G and Johnson GN (2004) Reduction of the
thylakoid electron transport chain by stromal reductants–evidence for activation of cyclic electron transport upon dark
adaptation or under drought. Planta 220: 356–363
Harbinson J (1994) The responses of thylakoid electron transport
and light utilization efficiency to sink limitation of photosynthesis. In: Baker NR and Bowyer JR (eds) Photoinhibition of Photosynthesis, from Molecular Mechanisms to the
Field, pp 273–295. BIOS scientific publishers Springer, The
Netherlands
Harbinson J and Foyer CH (1991) Relationships between the
efficiencies of Photosystem-I and Photosystem-II and stromal
redox state in CO2 -free air – evidence for cyclic electron flow
in vivo. Plant Physiol 97: 41–49
Harbinson J and Hedley CL (1993) Changes in P-700 oxidation
during the early stages of the induction of photosynthesis.
Plant Physiol 103: 649–660
Harbinson J, Genty B and Baker NR (1990) The relationship
between CO2 assimilation and electron transport in leaves.
Photosynth Res 25: 213–224
Hashimoto M, Endo T, Peltier G, Tasaka M and Shikanai T (2003)
A nucleus-encoded factor, CRR2, is essential for the expression of chloroplast ndhB in Arabidopsis. Plant J 36: 541–549
Hauser M, Eichelmann H, Oja V, Heber U and Laisk A (1995)
Stimulation by light of repid pH regulation in the chloroplast
stroma in vivo as indicated by CO2 solubilization in leaves.
Plant Physiol 108: 1059–1066
Heber U (2002) Irrungen, Wirrungen? The Mehler reaction in
relation to cyclic electron transport in C3 plants. Photosynth
Res 73: 223–231
Heber U and Walker D (1992) Concerning a dual function of
coupled cyclic electron-transport in leaves. Plant Physiol 100:
1621–1626
Heber U, Egneus H, Hanck U, Jensen M and Koster S (1978)
Regulation of photosynthetic electron transport and phosphorylation in intact chloroplasts and leaves of Spinacia oleracea
L. Planta 143: 41–49
Heber U, Neimanis S, Siebke K, Schonknecht G and Katona E
(1992) Chloroplast energization and oxidation of P700 and
plastocyanin in illuminated leaves at reduced levels of CO2 or
oxygen. Photosynth Res 34: 433–447
Herbert SK, Fork DC and Malkin R (1990) Photoacoustic measurements in vivo of energy storage by cyclic electron transport
in algae and higher plants. Plant Physiol 94: 926–934
Hill R and Bendall F (1960) Function of the two cytochrome components in chloroplasts: a working hypothesis. Nature 186:
136–137
Hiyama T and Ke B (1971) A further study of P430. A possible
primary acceptor of photosystem I. Arch Biochem Biophys
147: 99–180
Horvath EM, Peter SO, Joet T, Rumeau D, Cournac L, Horvath GV, Kavanagh TA, Schafer C, Peltier G and Medgyesy
P (2000) Targeted inactivation of the plastid ndhB gene in tobacco results in an enhanced sensitivity of photosynthesis to
moderate stomatal closure. Plant Physiol 123: 1337–1349
Pierre Joliot, Anne Joliot and Giles Johnson
Hosler JP and Yocum CF (1985) Evidence for 2 cyclic photophosphorylation reactions concurrent with ferredoxin-catalyzed
non-cyclic electron-transport. Biochim Biophys Acta 808: 21–
31
Jagendorf AT and Avron M (1958) Cofactors and rates of photosynthetic phosphorylation by spinach chloroplasts. J Biol
Chem 231: 277–290
Jeanjean R, Bedu S, Havaux M, Matthijs HCP and Joset F (1998)
Salt-induced photosystem I cyclic electron transfer restores
growth on low inorganic carbon in a type 1 NAD(P)H dehydrogenase deficient mutant of Synechocystis PCC 6803. FEMS
Microbiol Lett 167: 131–137
Joët T, Cournac L, Peltier G and Havaux M (2002) Cyclic electron
flow around photosystem I in C-3 plants. In vivo control by
the redox state of chloroplasts and involvement of the NADHdehydrogenase complex. Plant Physiol 128: 760–769
Johnson GN (2003) Thiol regulation of the thylakoid electron
transport chains: a missing link in the regulation of photosynthesis? Biochemistry 42: 3040–3044
Johnson GN (2005) Cyclic electron transport in C3 plants: fact
or artefact? J Exp Bot 56: 407–416
Joliot P and Joliot A (1988) The low-potential electron-transfer
chain in the cytochrome b/ f complex. Biochim Biophys Acta
933: 319–333
Joliot P and Joliot A (2002) Cyclic electron transfer in plant leaf.
Proc Natl Acad Sci USA 99: 10209–10214
Joliot P and Joliot A (2004) Cyclic electron flow under saturating excitation of dark-adapted Arabidopsis leaves. Biochim
Biophys Acta 1656: 166–176
Joliot, P and Joliot A (2005) Quantification of cyclic and linear
flows in plants. Proc Natl Acad Sci USA 102: 4913–4918
Joliot P, Lavergne J and Béal D (1992) Plastoquinone compartmentation in chloroplasts. 1. Evidence for domains with different rates of photo-reduction. Biochim Biophys Acta 1101:
1–12
Joliot P, Beal D and Joliot A (2004) Cyclic electron flow
under saturating excitation of dark-adapted Arabidopsis
leaves. Biochim Biophys Acta 1656: 166–176
Katona E, Neimanis S, Schonknecht G and Heber U (1992) Photosystem I-dependent cyclic electron-transport is important in
controlling Photosystem-II activity in leaves under conditions
of water-stress. Photosynth Res 34: 449–464
Kirchhoff H, Horstmann S and Weis E (2000) Control of the photosynthetic electron transport by PQ diffusion microdomaines
in thylakoids of higher plants. Biochim Biophys Acta 1459:
148–168
Klughammer C and Schreiber U (1994) An improved method,
using saturating light-pulses, for the determination of
Photosystem-I quantum yield via P+
700 - absorbency changes
at 830 nm. Planta 192: 261–268
Kurisu G, Zhang HM, Smith JL and Cramer WA (2003) Structure
of the cytochrome b6 f complex of oxygenic photosynthesis:
tuning the cavity. Science 302: 1009–1014
Laisk A (1993) Mathematical modelling of free-pool and channelled electron transport in photosynthesis: evidence for a
functional supercomplex around photosystem 1. Proc R Soc
Lond B 251: 243–251
Laisk A, Oja V and Heber U (1992) Steady-state and induction kinetics of the photosynthetic electron transport related to donor
side oxidation and acceptor side reduction of photosystem 1
in sunflower leaves. Photosynthetica 27: 449–463
Chapter 37
Cyclic Electron Transfer Around Photosystem I
Lascano HR, Casano LM, Martin M and Sabater B (2003) The
activity of the chloroplastic Ndh complex is regulated by phosphorylation of the NDH-F subunit. Plant Physiol 132: 256–262
Lavergne J (1983) Membrane potential-dependent reduction of
cyt b6 in algal mutant lacking Photosystem I centers. Biochim
Biophys Acta 725: 25–33
Lavergne J, Bouchaud JP and Joliot P (1992) Plastoquinone compartmentation in chloroplasts. 2. Theoretical aspects. Biochim
Biophys Acta 1101: 13–22
Matthijs HCP, Jeanjean R, Yeremenko N, Huisman J, Joset F
and Hellingwerf KJ (2002) Hypothesis: versatile function of
ferredoxin-NADP(+) reductase in cyanobacteria provides regulation for transient photosystem I-driven cyclic electron flow.
Funct. Plant Biol. 29: 201–210
Maxwell PC and Biggins J (1976) Role of cyclic electron transport in photosynthesis as measured by photoinduced turnover
of P700 in vivo. Biochemistry 15: 3975–3981
Mehler AT (1951) Studies on reactions of illuminated chloroplasts. I. Mechanism of the reduction of oxygen and other Hill
reagents. Arch Biochem Biophys 33: 65–77
Mi HL, Endo T, Schreiber U and Asada K (1992a) Donation of
electrons from cytosolic components to the intersystem chain
in the cyanobacterium Synechococcus sp PCC 7002 as determined by the reduction of P700+ . Plant Cell Physiol 33:
1099–1105
Mi HL, Endo T, Schreiber U, Ogawa T and Asada K (1992b)
Electron donation from cyclic and respiratory flows to the
photosynthetic intersystem chain is mediated by pyridinenucleotide dehydrogenase in the cyanobacterium Synechocystis PCC 6803. Plant Cell Physiol 33: 1233–1237
Mi HL, Klughammer C and Schreiber U (2000) Light-induced
dynamic changes of NADPH fluorescence in Synechocystis
PCC 6803 and its ndhB-defective mutant M55. Plant Cell
Physiol 41: 1129–1135
Miyake C, Shinzaki Y, Miyata M and Tomizawa K (2004) Enhancement of cyclic electron flow around PSI at high light and
its contribution to the induction of non-photochemical quenching of chl fluorescence in intact leaves of tobacco plants. Plant
Cell Physiol 45: 1426–1433
Miyake C, Horiguchi S, Makino A, Shinzaki Y, Yamamoto
H and Tomizawa K (2005a) Effects of light intensity on
cyclic electron flow around PS I and its relationship to nonphotochemical quenching of Chl fluorescence in tobacco
leaves. Plant Cell Physiol 46: 1819–1830
Miyake C, Miyata M, Shinzaki Y and Tomizawa K (2005b) CO2
response of cyclic electron flow around PS I (CEF-PS I) in
tobacco leaves–relative electron fluxes through PS I and PS II
determine the magnitude of non-photochemical quenching
(NPQ) of Chl fluorescence. Plant Cell Physiol 46: 629–637
Mitchell P (1975) The protonmotive Q cycle: a general formulation. FEBS Lett 59: 137–199
Moss DA and Bendall DS (1984) Cyclic electron transport in
chloroplasts. The Q-cycle and the site of action of antimycin.
Biochim Biophys Acta 767: 389–395
Mullineaux CW (1999) The thyalkoid membranes of cyanobacteria: structure dynamics and function. Aust J Plant Physiol
26: 671–677
Munekage Y, Hojo M, Meurer J, Endo T, Tasaka M and Shikanai
T (2002) PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in Arabidopsis.
Cell 110: 361–371
655
Ogawa T (1991) Cloning and inactivation of a gene essential to
inorganic carbon transport of Synechocystis PCC 6803. Plant
Physiol 96: 280–284
Oxborough K and Horton P (1987) Characterisation of the effects
of antimycin A upon high-energy-state quenching of chlorophyll fluorescence (qE) in spinach and pea chloroplasts. Photosynth Res 12: 119–128
Sacksteder A and Kramer DM (2000) Dark-interval relaxation
kinetics (DIRK) of absorbance changes as a quantitative
probe of steady-state electron transfer. Photosynth Res 66:
145–158
Sandmann G and Malkin R (1983) NADH and NADPH as electron donors to respiratory and photosynthetic electron transport in the blue-green alga Aphanocapsa. Biochim Biophys
Acta 725: 21–224
Sandmann G and Malkin R (1984) Light inhibition of respiration is due to a dual function of the cytochrome b6 f complex
and the plastocyanin/cytochrome c-533 pool in Aphanocapsa.
Arch Biochem Biophys 234: 105–111
Sazanov LA, Burrows P and Nixon PJ (1995) Presence of a large
protein complex containing the ndhK gene product and possessing NADH-specific dehydrogenase activity in thylakoid
membranes of higher plant chloroplasts. In: Mathis P (ed)
Photosynthesis: From Light to Biosphere Xth International
Congress on Photosynthesis, Vol 2, pp 705–708. Kluwer Academic Publishers, Montpellier, France
Sazanov LA, Burrows PA and Nixon PJ (1998) The chloroplast
Ndh complex mediates the dark reduction of the plastoquinone
pool in response to heat stress in tobacco leaves. FEBS Lett
429: 115–118
Schankser G, Srivastava A, Govindjee and Strasser RJ (2003)
Characterisation of the 820-nm transmission signal paralleling
the chlorophyll a fluorescence rise (OIJP) in pea leaves. Funct
Plant Biol 30: 785–796
Scheller H (1996) In vitro cyclic electron transport in barley
thylakoids follows two independent pathways. Plant Physiol
110: 187–194
Scherer S (1990) Do photosynthetic and respiratory electron
transport chains share redox proteins? TIBS 15: 458–462
Schmetteter G (1994) Cyanobacterial respiration. In: Bryant DA
(ed) The Molecular Biology of Cyanobacteria, Vol 1, pp 409–
435. Kluwer Academic Publishers, Dordrecht
Seelert H, Poetsch A, Dencher NA, Engel A, Stahlberg H and
Muller DJ (2000) Structural biology. Proton-powered turbine
of a plant motor. Nature 405: 418–419
Sherman DM, Troyan TA and Sherman LA (1994) Localization
of membrane proteins in the cyanobacterium Synechococcus
sp. PCC 7942. Plant Physiol 106: 251–262
Shinokazi K, Ohme M, Tanaka M, Wakasuki T, Hayashida
N, Matsubayashi T, Zaita N, Chunwongse J, Obokata J,
Yamaguchi-Shinokazi K, Ohto C, Torazawa K, Meng BY,
Sugita M, Deno H, Kamogashira T, Yamada K, Kusuda J,
Takaiwa F, Kato A, Tohdoh N, Shimada H and Sugiura M
(1986) The complete nucleotide sequence of the tobacco
chloroplast genome: its gene organization and expression.
EMBO J 5: 2043–2049
Smith L and Baltscheffsky M (1959) Respiration and lightinduced phosphorylation in extracts of Rhodospirillum
rubrum. J Biol Chem 234: 1575–1579
Strasser RJ, Schansker G, Srivastava A and Govindjee (2001) Simultaneous measurement of Photosystem I and Photosystem
656
II probed by modulated transmission at 820 nm and by chlorophyll a fluorescence in the sub ms to second time range.
In: Critchley C (ed) PS2001 12th International Congress on
Photosynthesis, pp S14–003. CSIRO Publishers, Brisbane,
Australia
Stroebel D, Choquet Y, Popot J-L and Picot D (2003) An atypical haem in the cytochrome b6 f complex. Nature 426: 413–
418
Tagawa K, Tsujimoto HY and Arnon DI (1963a) Analysis of
photosynthetic reactions by the use of monochromatic light.
Nature 199: 1247–1252
Tagawa K, Tsujimoto HY and Arnon DI (1963b) Role of chloroplast ferredoxin in the energy conversion process of photosynthesis. Proc Natl Acad Sci USA 49: 567–572
Teicher BH, Moller BL and Scheller HV (2000) Photoinhibition
of Photosystem I in field-grown barley (Hordeum vulgare L.):
induction, recovery and acclimation. Photosynth Res 64: 53–
61
Vallon O, Bulte L, Dainese P, Olive J, Bassi R and Wollman FA
(1991) Lateral redistribution of cytochrome b6 / f complexes
along thylakoid membranes upon state transitions. Proc Natl
Acad Sci USA 88: 8262–8266
van Thor JJ, Jeanjean R, Havaux M, Sjollema KA, Joset F,
Hellingwerf KJ and Matthijs HCP (2000) Salt shock-inducible
Photosystem I cyclic electron transfer in Synechocystis PCC
6803 relies on binding of ferredoxin:NADP(+) reductase to
the thylakoid membranes via its CpcD phycobilisome-linker
homologous N-terminal domain. Biochim Biophys Acta 1457:
129–144
Pierre Joliot, Anne Joliot and Giles Johnson
Wang HW, Mi HL, Ye JY, Deng Y and Shen YK (2003) Low
concentrations of NaHSO3 increase cyclic photophosphorylation and photosynthesis in cyanobacteriumSynechocystis PCC
6803. Photosynth Res 75: 151–159
Webber AN, Platt-Aloia KA, Heath RL and Thomson WW
(1988) The marginal regions of thylakoid membranes: a partial characterization by polyoxyethylene sorbitane monolaurate (Tween 20) solubilization of spinach thylakoids. Physiol
Plant 72: 288–297
Whatley FR (1963) Some effects of oxygen in photosynthesis by
chloroplast preparations. In: Kok B and Jagendorf A (eds)
Photosynthetic Mechanisms of Green Plants, pp 243–250.
National Academy of Sciences – National Research Council,
Washington, DC
Whatley FR, Allen MB and Arnon DI (1959) Photosynthesis
by isolated chloroplasts. VII. Vitamin K and Riboflavin Phosphate as cofactors of cyclic photophosphorylation. Biochim
Biophys Acta 32: 32–46
Wollman F-A and Bulté L (1989) Toward an understanding of the
physiological role of state transitions. In: Hall DO and Grassi
G (eds) Photoconversion Processes for Energy and Chemicals,
pp 198–207. Elsevier, Amsterdam
Yu L, Zhao JD, Muhlenhoff U, Bryant DA and Golbeck JH
(1993) PsaE is required for in vivo cyclic electron flow around
Photosystem-I in the cyanobacterium Synechococcus sp. PCC7002. Plant Physiol 103: 171–180
Zhang HM, Whitelegge JP and Cramer WA (2001) Ferredoxin:NADP(+) oxidoreductase is a subunit of the chloroplast
cytochrome b(6)f complex. J Biol Chem 276: 38159–38165