cyclic electron transfer around photosystem i€¦ · cyclic electron transport around photosystem...

Chapter 37

Cyclic Electron Transfer Around Photosystem I

Pierre Joliot∗ and Anne JoliotCNRS UMR 7141, Institut de Biologie Physico-Chimique, 13, rue Pierre et Marie Curie, 75005

Paris, France

Giles JohnsonUniversity of Manchester, School of Biological Sciences, 3.614 Stopford Building, Oxford Road,

Manchester, M13 9PT, UK

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 639I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640II. Early Observations of Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640III. Possible Pathways of Electron Flow in Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640IV. Redox Poising of the Cyclic Electron Transfer Chain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641V. Structural Organization of Thylakoid Membranes – Consequences for Cyclic Electron Transfer . . . . . . . . . . . 642VI. Occurrence of Cyclic Flow in Higher Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643VII. Pathway of Cyclic Flow in Higher Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647

A. NADPH-Dependent Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647B. Ferredoxin-Dependent Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647

VIII. Cyclic Flow in Green Unicellular Algae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649IX. Cyclic Flow in Cyanobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650X. Functions and Regulation of Cyclic Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651XI. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .652Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 652References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 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 physiologicalrole and even its very occurrence remain in question. Nevertheless, significant progress is starting to be made in ourunderstanding of this process. At least two pathways of cyclic electron transport appear to operate, one involvingthe transfer of electrons from NADPH to plastoquinone and the other operating via the donation of electrons fromferredoxin 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 itsfunctional organization in the thylakoid membrane. Two distinct functions for cyclic electron transport can bedefined — the generation of ATP and, in higher plants, the generation of �pH to regulate light harvesting. Thesetwo functions give rise to the need for different regulatory processes to control the ratio of cyclic and linear electronflow. We discuss recent findings that cast new light on how cyclic electron transport is regulated under a range ofphysiological conditions.

∗Author for correspondence, email: [email protected]

John H. Golbeck (ed): Photosystem I: The Light-Driven Plastocyanin:Ferredoxin Oxidoreductase, 639–656.C© 2006 Springer.

640 Pierre Joliot, Anne Joliot and Giles Johnson

I. Introduction

The concept of cyclic electron transport (ET) aroundPhotosystem (PS) I is sufficiently established that it fea-tures in the diagrams of photosynthetic electron trans-port found in every undergraduate biochemistry text-book. In spite of this celebrity, the redox componentsinvolved, the functional importance, the regulation and,indeed, even the very occurrence of this pathway all re-main unclear. Part of the problem in studying cyclic EThas been, that, by its very nature, it is difficult to quan-tify — as a cycle it involves no net flux and so we areforced to resort to use indirect means to deduce its ex-istence. There is now growing evidence, however, thatcyclic ET does indeed occur in a variety of organismsunder a wide range of conditions, fulfilling at least twodistinct functions. Here, we present a review of thisevidence, focusing in particular on advances over thelast decade and identifying the challenges that remain.For more detailed coverage of earlier work and alterna-tive views on the subject, a number of good reviews areavailable (Heber and Walker, 1992; Fork and Herbert,1993; Bendall and Manasse, 1995; Heber, 2002; Allen,2003).

II. Early Observations of Cyclic ElectronTransfer

Photosynthetic phosphorylation (photophosphoryla-tion) was discovered in 1954 by Arnon et al. (1954) whoestablished that illumination of isolated chloroplasts inthe presence of oxygen-induced ATP synthesis. As thisprocess was not associated with oxygen formation orconsumption, the contribution of a respiratory chaincould be excluded. Later, it was established that the de-pendence of photophosphorylation on oxygen can beabolished by the addition of vitamin K or other naph-thoquinones (Arnon, 1955). This experiment marks thediscovery of the cyclic phosphorylation process, (re-viewed in Arnon et al., 1961). The reaction previouslycharacterized in 1954 by Arnon and coworkers appearsnow to be a pseudocyclic process in which electronsare transferred from water to O2 (Mehler, 1951) via the

Abbreviations: cyt – cytochrome; DBMIB – 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone; DCMU – 3-(3,4-dichloro-phenyl)-1,1-dimethylurea; ET – electron transport;ETC – electron transfer chain; Fd – ferredoxin; FNR – ferre-doxin: NADP oxidoreductase; FQR – ferredoxin: plastoquinonereductase; HQNO – 2-heptyl-4-hydroxy-quinoline N-oxide;NDH – NADH dehydrogenase; PC – plastocyanin; PMS – N-methylphenazonium-3-sulfonate; PQ – plastoquinone; PQH2 –plastoquinol; PS – Photosystem; RC – reaction center.

linear electron transfer chain. Subsequently, nonphys-iological compounds, such as phenazine methosulfate(PMS), were shown to catalyze cyclic photophospho-rylation more efficiently than vitamin K (Jagendorf andAvron, 1958).

The anaerobic cyclic phosphorylation identified inchloroplasts appeared analogous to a cyclic phosphory-lation process previously identified in chromatophoresfrom photosynthetic bacteria (Frenkel, 1954). In Rho-dospirillum rubrum, this cyclic process was observedto involve cytochrome (cyt) c (Smith and Baltscheff-sky, 1959); in chloroplasts cyt f is involved (Arnon,1959). In 1960, Hill and Bendall put forward a modeldescribing the linear electron transfer chain involvingboth the photoreactions PS II and PS I working in se-ries (Hill and Bendall, 1960). It became clear, however,that cyclic photophosphorylation involves only PS I,as it operates in the presence of specific inhibitors ofoxygen evolution, such as 3-(3,4-dichloro-phenyl)-1,1-dimethylurea (DCMU). Tagawa et al. (1963b) estab-lished that ferredoxin (Fd) was able to catalyze cyclicphosphorylation. This provided evidence for a physi-ological catalyst of cyclic ET, making it seems likelythat this process was more than an artifact of in vitroconditions.

III. Possible Pathways of Electron Flowin Cyclic Electron Transfer

There is a general agreement that both PS I and the cytb/ f complex are obligatory involved in cyclic elec-tron flow — cyclic ET is efficiently driven by far-red light and is sensitive to inhibitors of the cyt b/ fcomplex, such as 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) and stigmatellin — how-ever, the electron pathway from the acceptor side of PSI back to the cyt b/ f complex has not yet been clearlyelucidated. Cyclic ET, as conventionally measured,leads to the generation of a trans-thylakoid pH gra-dient. Thus, whatever the pathway involved a proton-pumping step involving the cyt b/ f complex must bepostulated.

Based mainly on studies of the effects of the in-hibitor antimycin A, it has been postulated that at leasttwo pathways exist. Experiments performed on isolatedchloroplasts showed that antimycin inhibits cyclic flowin the presence of Fd (Tagawa et al., 1963b) but notin the presence of vitamin K or PMS (Whatley et al.,1959). Hosler and Yocum (1985) measured the ratio ofP/O in the presence of Fd, using oxygen as a terminalelectron acceptor and found this to be sensitive to an-timycin. 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 ex-periment was performed using NADP as an electron ac-ceptor, the P/O ratio was high and antimycin-insensitive(Hosler and Yocum, 1985). Scheller (1996) measuredthe rereduction of P700 following light flashes in thepresence of DCMU and noted an antimycin-insensitiveportion in Fd-mediated cyclic ET, possibly suggesting athird pathway (Scheller, 1996). However, the very slowrate of this, barely distinguishable from the DBMIB-inhibited rate, makes it hard to separate this from back-ground redox equilibration of the sample. Cleland andBendall (1992) measured the oxidation of reduced Fdfollowing illumination of DCMU-poisoned thylakoidsand found this rate to be inhibited by antimycin A ascompletely as by stigmatellin (Cleland and Bendall,1992). One can thus conclude that several mechanismscould be involved in a cyclic ET.

In a conventional Q-cycle process (Mitchell, 1975;Crofts et al., 1983), electrons are transferred to the cytb/ f complex via a reduced quinone that binds site Qo,on the lumenal side of this complex. Assuming thisis the step generating �pH in the cyclic process, weneed to invoke an enzyme that is able to transfer elec-trons from NADP or Fd to plastoquinone (PQ), withthe resultant plastoquinol (PQH2) being protonatedon the stromal side of the membrane. In cyanobacteria,the presence of respiratory and photosynthetic electrontransport chains in the same membrane means that thisfunction could be fulfilled by a respiratory NADH de-hydrogenase (NDH; complex I). Genes coding for suchan enzyme have also been identified in the chloroplastgenome of higher plants (Shinokazi et al., 1986), which

is a likely candidate to be involved in the cyclic electrontransfer chain (ETC) (Fig. 1, pathway 1).

Moss and Bendall (1984) noted that, whilst both Fd-mediated and artificially mediated cyclic ET are sensi-tive to the cyt b inhibitor 2-heptyl-4-hydroxy-quinolineN-oxide (HQNO), only Fd-mediated cyclic ET wassensitive to antimycin (Moss and Bendall, 1984). Thisled them to suggest an alternative site for antimycininhibition on an enzyme distinct from the b/ f com-plex, termed ferredoxin:plastoquinone oxidoreductase(FQR) (Fig. 1, pathway 2).

The involvement of ferredoxin:NADP oxidoreduc-tase (FNR) in cyclic ET has been much discussed. Itsinvolvement in the NADP-dependent pathway is pre-sumed; however, a role in the Fd-dependent pathwayhas also been postulated. The observation that this en-zyme is stoichiometrically bound to the cyt b/ f com-plex in spinach provides an intriguing indication of anadditional role, other than that of linear electron trans-port (Zhang et al., 2001). It is suggested that FNR me-diates the transfer of electrons from ferredoxin to siteQi (Fig. 1, pathway 3).

The mechanisms involved in these different path-ways will be discussed in more detail below.

IV. Redox Poising of the Cyclic ElectronTransfer Chain

Photochemical charge separation at the level of areaction center (RC) requires the presence of a re-duced primary donor and an oxidized electron acceptor.


Over-reduction or overoxidation of the cyclic chain willthus inhibit this process. The concept of redox poisingof the cyclic ETC was introduced to explain the oxygenrequirement of cyclic phosphorylation (Tagawa et al.,1963a; Whatley, 1963; reviewed in Allen, 1983). Later,Arnon and Chain (1977) established that a maximumefficiency of the cyclic ET in the presence of oxygenis observed in the presence of NADPH and subsaturat-ing concentrations of DCMU that impose an optimalredox poise of the carriers involved in the cyclic chain(Arnon and Chain, 1977).

Redox poising is determined by the relative rate ofelectron efflux or influx from or toward the carriersbelonging to the cyclic electron transfer chain. Bothefflux and influx will occur preferentially at the levelof mobile carriers that are common to linear and cyclicchains. PS II will be the main source of reductive power(Fig. 1, pathways a and b) while electron efflux willoccur at the level of PS I acceptors toward O2 and theBenson–Calvin cycle (Fig. 1, pathways c and d, respec-tively). If we assume that cyclic and linear pathways areconnected, i.e., share the same mobile carriers, the re-dox poise of the cyclic chain will be controlled by theelectron flow through the linear chain. Under strong il-lumination, given under condition where the Benson–Calvin cycle is inhibited (e.g., in isolated thylakoids ordark-adapted leaves), PS II will induce an overreduc-tion of the cyclic chain, via pathway a or b (Fig. 1).Conversely, under conditions where PS II is inhibitedor under far-red light, the electron efflux via pathway cor d will induce an overoxidation of the cyclic chain. Onthe other hand, if the cyclic and linear chains are struc-turally separated, the redox poise of the cyclic chainwill be controlled by the rate of slow electron leaksthat occur between carriers involved in both processes.We thus conclude that the structural organization ofmembrane proteins that controls the localization of thecyclic and linear chains within the membrane may playan essential role in the control of the efficiency of thecyclic process.

V. Structural Organization of ThylakoidMembranes – Consequences forCyclic Electron Transfer

Oxygenic photosynthetic organisms — higher plants,unicellular algae, and cyanobacteria — differ substan-tially in the supramolecular organization of their pho-tosynthetic membranes and these differences may haveimportant consequences for the pathway and regulationof cyclic ET.

In higher plants, PS II and PS I are localized in dif-ferent membrane regions (Andersson and Anderson,1980). Most of PS II is localized in the appressedregions of the grana stacks while PS I is localizedin the stroma lamellae and in the granal end mem-branes. Unlike PS II and PS I, cyt b/ f complex isdistributed across all membrane regions (Cox and An-dersson, 1981). An open question concerns a possi-ble localization of PS I centers in the margin of thegrana stacks (Webber et al., 1988; Anderson, 1989;Albertsson, 1995). Albertsson (2001) put forward thehypothesis that PS I localized in the margin and endsof the grana stacks contributes to the linear pathwaywhile PS I localized in the stroma lamellae contributesto the cyclic pathway (Albertsson, 2001). The local-ization of PS I and PS II in different membrane re-gions requires long-range diffusion of the mobile car-riers PQ or plastocyanin (PC). A detailed analysis ofthe kinetics of electron transfer reaction between PS IIand the PQ pool has shown that diffusion of PQ is re-stricted to small heterogeneous domains including anaverage of three to four RC, a membrane surface muchsmaller than the size of a grana disk (Joliot et al., 1992;Lavergne et al., 1992; Kirchhoff et al., 2000). It is as-sumed that the membrane proteins, which occupy morethan half of the membrane surface, limit the diffusionof PQ. This implies that PQ is not involved in long dis-tance transfer and that the linear process exclusivelyinvolves cyt b/ f complexes localized close to PS II,i.e., in the appressed regions. Conversely, the sole roleof the cyt b/ f complexes localized in the stroma regionmight be to participate in the cyclic process (“cyclic cytb/ f ”).

It worth pointing out that, if the photosynthetic ap-paratus were exclusively devoted to a linear process,one would expect that its optimization during evolu-tion would have led to a random distribution of RCswithin the membrane. Such a distribution would mini-mize the distance between membrane proteins, leadingto faster electron exchanges mediated by PQ or PC.Thus, the segregation of PS I and PS II centers in differ-ent membrane regions can be taken as a way to separatethe carriers involved in the cyclic and the linear flows,which limits redox cross-talk between the cyclic andlinear chains. Structural separation between the linearand cyclic processes is pushed to an extreme in the caseof C4 plants, in which only a cyclic process operatesin bundle sheath cells that mainly include PS I centers(Bassi et al., 1985).

In green unicellular algae, the supramolecular or-ganization of thylakoid membranes significantly dif-fers from that in higher plants. Thylakoid membranes


consist of long flat vesicles (disks) that are generallystacked in groups of 2–4, a much smaller number thanthat seen in higher plants. Freeze-fracture images sug-gest that appressed and nonappressed regions are morewidely connected than is the case in chloroplasts ofhigher plants, which are connected by narrow fret junc-tions. This membrane organization suggests that cyclicand linear pathways could interact more in green algaethan in higher plants.

In cyanobacteria, thylakoids membranes are un-stacked and appear as isolated flat vesicles. The orga-nization of these membranes differs between speciesbut concentric arrangements of thylakoids are of-ten seen in rod-shaped cells as Synechococcus sp.PCC 6803 or filamentous species such as Phormidiumlaminosum. There is evidence from freeze-fracture im-ages of cyanobacterial thylakoid membranes that PS Iand PS II are typically found in the same membraneregions — see Mullineaux (1999), although Shermanet al. (1994) noted a slight asymmetry in the distri-bution of complexes in Synechococcus sp. PCC 6803,suggesting a concentration of PS I near to the plasmamembranes. Spatial segregation of linear and cyclicchains is thus very unlikely and one expects these path-ways to share the same electron carriers. In contrastto photosynthetic eukaryotes, thylakoids in cyanobac-teria include a respiratory chain that shares the PQpool and the cyt b/ f complex with the photosyntheticchain (for a review see Schmetteter, 1994). Thus, re-dox poising of a putative cyclic chain could be con-trolled by interaction with the linear photosyntheticchain as well as with the respiratory chain. An openquestion is the nature of the substrate of the NDH en-zymes of the respiratory chain—NADH or NADPH—localized in the thylakoids. NADPH-PQ reductase,present at high concentration in the chloroplasts, coulditself contribute efficiently to a cyclic process (Fig. 1,pathway 1).

VI. Occurrence of Cyclic Flow inHigher Plants

Work establishing the existence of pathways for cyclicET has largely been performed in isolated systems, usu-ally thylakoid membranes (broken chloroplasts), withaddition of natural or artificial mediators. While suchstudies are essential in characterizing the pathway ofcyclic ET, they leave open the question; does this path-way actually operate under in vivo conditions? Morespecifically, if we are to understand the function ofcyclic ET it must be established whether it occurs un-

der normal physiological conditions, where linear ETis also possible.

Many of the studies that have indicated the presenceof cyclic ET in intact leaves have used rather indirectmeans, typically involving conditions that largely ortotally suppress PS II turnover. While such studies arevaluable, especially in trying to understand the regula-tion of the cyclic pathway, they do not, in themselves,show that this pathway is able to compete with linearET.

One commonly used assay taken as evidence forcyclic flux is the measurement of the relaxation ofP700+ following illumination of a leaf with a periodof far-red light (� > 695 nm) or in the presence of aPS II inhibitor, such as DCMU (Maxwell and Biggins,1976). When a leaf is exposed to far-red light to oxi-dize P700 and then that light is abruptly cut, typicallyat least two phases of P700+ reduction can be detected.The half time for the fast phase is typically of the orderof 200–1,000 msec (Joet et al., 2002). By comparison,in white light, under conditions where PS II is turningover normally, the half time for P700+ reduction is ofthe order of 10–20 msec. Thus, it seems immediatelyunlikely that cyclic ET could ever compete effectivelywith a linear flow that is 10–50 times faster. However,measurements made in the absence of PS II turnoverwill tend to result the accumulation of electron trans-fer components (including Fd and NADP) in the ox-idized state and so, the half times measured probablyrepresent a gross underestimate of the maximum rateof electron flow from the stroma to P700+. The rateof P700+ reduction varies between different groups oforganisms, being slow in plants and especially slow inC3 plants (Herbert et al., 1990; Joet et al., 2002). Therate can, however, be accelerated under certain condi-tions, for example under anaerobiosis (Joet et al., 2002)or following heat stress (Maxwell and Biggins, 1976;Burrows et al., 1998; Bukhov et al., 1999). The formerprobably reflects an increase in the reduction state of thechloroplast in the dark, increasing the supply of reduc-tant to re-reduce P700+. In the latter case, it is less clearwhat gives rise to the effect, it may relate to a tempera-ture induced shift in redox poise or to an “opening up”of redox components to the surrounding medium accel-erating their reduction. Recently, Golding et al. (2004),examining the relaxation of P700+ in the presence ofthe PS II inhibitor DCMU, observed that preillumina-tion of leaves eliminates the fast component of P700+

reduction but that this effect is removed if the leavesare experiencing drought stress. It is suggested that atransient “cyclic-enabled” state existing in the dark isstabilized under drought conditions (Johnson, 2005).


Another parameter that has been taken to indicatecyclic ET is the presence of a transient rise in fluo-rescence 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 seento rise transiently and then fall again. This effect is at-tributed to the reduction of the PQ pool by electronsoriginating from the stroma, indicating that a pathwayexists between the stroma and the electron transportchain that could participate in cyclic ET. Such measure-ments can be made following conditions of steady-statewhite light, so may reflect better the normal physiolog-ical state of the leaf but, as with the decay of P700+,involve processes that are far too slow (several tens ofseconds) for them to be supposed to be involved in effi-cient cyclic ET in competition with linear ET. As withthe decay of P700+ following far-red light, this effectis enhanced by exposing plants to heat stress (Sazanovet al., 1998).

Although cyclic ET involves no net flux that canbe measured, it does nonetheless have “products” thatare measurable, notably the formation of a pH gradi-ent across the thylakoid membrane, which can be ev-idenced using optical spectroscopy and the storage oflight energy, which is measured using the technique ofphotoacoustics.

Two absorbance signals have been used as indicatorsof the pumping of protons across the thylakoid mem-brane during cyclic ET: Proton translocation resultsin the formation of a transmembrane electrical poten-tial, which can be measured as an apparent absorbancechange around 515 nm; and the swelling of the thy-lakoid membrane that occurs when a �pH is generatedinduces a change in its light scattering properties, giv-ing an apparent absorbance change in the region of535 nm. Both of these absorbance changes can be ob-served when leaves are illuminated with far-red light,indicating that cyclic ET is occurring and is generating�pH. However, the conditions needed to observe suchchanges are often quite specific, so again it is difficultto be certain whether the cycling observed is relevantto conditions of steady-state photosynthesis.

Photoacoustics measures pressure waves producedin samples in response to light, which can be inter-preted to provide information on energy storage andgas exchange. Photoacoustic signals are complex, witha number of different processes contributing, so it isnecessary to design experiments carefully to give anyinformation on cyclic ET. As with most other methods,this often means using far-red light to avoid any con-tribution from PS II photochemistry. A large number

of studies have been published using this approach, of-ten in combination with other approaches. For example(Joet et al., 2002), recently combined measurements ofphotoacoustics with relaxation of P700+ to investigatethe functioning of cyclic ET in tobacco.

In spite of the array of different methods that havebeen applied in an attempt to determine whether cyclicET occurs in higher plants, the question still remainscontroversial. A clear case where cyclic ET is thoughtto be the norm is in the bundle sheath cells of certainC4 plants, including maize (Herbert et al., 1990; Asadaet al., 1993; Joet et al., 2002). In such cells, PS II islargely absent, yet these cells are responsible for thefixation of CO2 through normal Benson–Calvin cycleactivity. By contrast, photosynthesis (and specificallyPS II) is not responsible for generating the reducingpotential required to drive CO2 fixation, since this isgenerated through the oxidation and decarboxylationof malate imported from the mesophyll. Cyclic ET isthought to provide the ATP. Studies using photoacous-tics and P700+ reduction kinetics have both providedsupport for the occurrence of cyclic ET in maize bun-dle sheath chloroplasts (Herbert et al., 1990; Joet et al.,2002) as have combined measurements of P700 andchlorophyll fluorescence under combinations of far-redand red light (Asada et al., 1993).

In C3 plants, the situation is less clear. As early as1978, Heber et al. (1978) observed the presence of farred-induced light scattering changes in spinach leaves.This scattering was slow to form however and was in-hibited by oxygen. The latter observation can easily beexplained in terms of the ability of oxygen to oxidizethe acceptor side of PS I. If PS II activity is suppressed,then electrons taking part in the cyclic pathway that areleaked to oxygen cannot be replaced. The cyclic ETCwill rapidly become completely oxidized and cyclic ETwill stop. This contrasts with the situation in C4 bundlesheath cells, where reductant is available from the de-carboxylation of malate, such that, even in the absenceof PS II activity, electrons can be reinjected in to thecyclic pathway.

A number of studies have, however, indicated that ef-ficient 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, underconditions of CO2 free air, far-red light was able to effi-ciently energize chloroplasts in cabbage leaves, as indi-cated by light scattering changes. This effect was how-ever suppressed by O2 or CO2. Heber et al. (1992) per-formed similar experiments on ivy leaves and reachedsimilar conclusions, with the CO2 concentration againbeing crucial. By contrast, the combination of red light


(rather than far-red) and low CO2 and O2 did not resultin chloroplast energization. In other words, conditionsgiving rise to PS II turnover in the absence of any termi-nal electron acceptor result in the rapid total reductionof the ETC. Under such conditions, no further electrontransport, linear or cyclic, is possible (see section onredox poising above). Taking an alternative approach,Cornic and colleagues observed the effects of periodsof actinic illumination on the ability of limiting levelsof far-red light to oxidize P700. Even short periods ofhigh light were found to be sufficient to reduce the ef-fectiveness of far-red light in oxidizing P700. This wasinterpreted as being due to an activation of cyclic ETduring high light, feeding electrons back into the ETCvia an efficient cyclic pathway (Cornic et al., 2000).

While the above discussion leads to the conclusionthat cyclic ET can be induced in the leaves of higherplants, it does not tell us whether it does occur un-der conditions of steady-state photosynthesis. Giventhe low rates of ET that have been measured and thecareful poising that is often needed to observe cyclicET, it is not at all clear that this pathway can competeunder conditions where linear ET is feeding electronsinto the ETC. To determine whether or not cyclic is areal physiological phenomenon, we need to be able tomeasure it under conditions of normal photosynthesis.

The most common approach evidencing cyclic ETunder conditions of steady-state photosynthesis is toexamine the relationship between PS I and PS II elec-tron transport. Given that cyclic ET involves only PSI and linear both PS I and PS II, any change in cyclicET, relative to linear ET will give a change in the ratioof the flux of the two photosystems. Measurements ofPS II flux are usually made using analysis of chloro-phyll fluorescence. The quantum efficiency of PS II ismeasured as the parameter �PS II (Genty et al., 1989).Multiplying this parameter by the light intensity givesa measure of relative flux. Provided light interceptionby the PS II antenna remains constant (which might notbe the case, e.g., due to state transitions) this parame-ter is thought to give a robust relative measure of thePS II electron transport rate.

In vivo measurements of PS I electron transportunder conditions of steady-state photosynthesis haveproved more controversial. A commonly used approachhas been to measure the redox state of the P700 pooland to take the extent of reduction of this pool as a mea-sure of the quantum efficiency of PS I. Comparisons ofPS I turnover measured in this way with PS II turnovermeasured by fluorescence have been made at a vari-ety of irradiances and CO2 concentrations (Harbinsonand Foyer, 1991; Harbinson, 1994). These studies have

found the relationship between these two parametersto be linear. Thus, it has been concluded that, undermost physiological conditions, cyclic ET is either ab-sent in the presence of light or forms a constant propor-tion of the linear flux. More recent data have howeverfound clear evidence for cyclic electron transport usingthis approach (Clark and Johnson, 2001; Miyake et al.,2004; Miyake et al., 2005a,b).

The contradictions between the above studies havenot yet been fully explained however probably relate tothe measuring conditions or the physiological status ofthe plants concerned. Observations of cyclic ET havebeen made under conditions where the supply of CO2 tothe leaf is restricted. For example, Harbinson and Foyer(1991) observed that the relationship between PS I andPS II quantum efficiency (using light as a variable) wasdifferent in CO2 free air to that seen in the presenceof CO2. Gerst et al. (1995) observed that, upon im-posing drought stress upon a leaf in the light, PS IIwas more sensitive to inhibition than PS I. By contrast,however, in experiments where a leaf was exposed tovarying CO2, the relationship between PS I and PS IIefficiency was found to be linear, extrapolating to theorigin (Harbinson, 1994). Thus any cyclic flow that oc-curs must be in proportion to linear flow. In a similarexperiment, Golding and Johnson (2003) noted that,although the total amount of reduced P700 was pro-portional to PS II efficiency, this relationship did notextrapolate to the origin, giving space for a constantrate of cyclic ET (Golding and Johnson, 2003). In ad-dition, however, these authors applied the method ofKlughammer and Schreiber (1994), to estimate the pro-portion of PS I in an “active” state. This measurementis performed by superimposing a saturating flash ofwhite light on top of background actinic illuminationand then transferring the sample directly to darkness,taking the total signal following the flash-to-dark tran-sition as a measure of active centers. In the study ofKlughammer and Schreiber (1994), the loss of activecenters was supposed to be related to a limitation on theacceptor side of PS I, preventing centers from turningover. Surprisingly, Golding and Johnson (2003) notedthat active PS I rose at low CO2 (i.e., under conditionswhere PS I is most likely to be acceptor-side limited).They suggested that a distinct population of PS I cen-ters exists that is largely or wholly involved in cyclicET. This “cyclic-only” pool is activated at low concen-trations of CO2 and under high light. Thus, the con-tradictions in earlier measurements might be relatedto the way in which CO2 limitation was applied andwhether these “cyclic” centers were already activatedor not.


In measurements of cyclic ET in far-red light it hasbeen common to take the rate of P700+ reduction asan indicator of PS I turnover (Maxwell and Biggins,1976). Essentially, the same approach can be appliedin white light conditions. This relies on the limiting stepin electron transport lying prior to PS I, which is usu-ally the case under physiological conditions, such thatthe flow of electrons to P700 can be measured and re-flects the overall flux through PS I. A potential problemwith this method was noted by Sacksteder and Kramer(2000), who noted that a net reduction of cyt f , mea-sured at the time the light is switched off, might haveto be taken into account to determine the electron fluxtoward P700 (Sacksteder and Kramer, 2000). Strictly,there should be no net reduction of cyt f at the momentwhen the light is switched off, as at steady-state the rateof oxidation and reduction are identical and neither ofthese are instantly affected by cutting the light. Practi-cally, given the time resolution and sensitivity of mostinstruments, this may be a problem but only under lowlight conditions where P700 is largely reduced but cytf oxidized, giving rise to a short time lag in the re-duction of cyt f . Sacksteder and Kramer (2000) havecompared the turnover of PS I and PS II in greenhousegrown sunflowers. They observed a linear relationshipbetween PS I and PS II ET, implying no (or a con-stant proportion of) cyclic ET, the same conclusion asreached by Harbinson and colleagues using P700 re-dox state to measure PS I turnover (Harbinson et al.,1990; Harbinson, 1994). In contrast, Clarke and John-son compared the rates of PS I and PS II turnover acrossa range of temperatures and light intensities in barleygrown in a growth cabinet. They observed that PS IIphotochemistry saturated at lower irradiances and wasmore sensitive to low temperature than PS I. Thus, theyconcluded that high light and low temperatures lead toenhancement of cyclic ET (Clarke and Johnson, 2001).The contradiction between these two studies may re-flect a species difference, but is more likely explainedby the range of light intensities used in each case. Inthe experiments of Sacksteder and Kramer, the high-est light intensities used were just saturating, whereasfor Clarke and Johnson light intensities were used thatwent well above saturating for PS II electron transport(though not necessarily for PS I). Thus it appears thatcyclic ET is a characteristic of saturating light, althougha low rate at subsaturating light cannot be excluded, ifthis forms a constant proportion of the linear flux. Asimilar approach by Golding and Johnson (2003) drewthe same conclusion concerning responses to low CO2

and drought.A new technical approach, based on membrane po-

tential measurements (Joliot and Joliot, 2002, 2004)

has been developed to determine the absolute rateof the cyclic and linear pathways whatever the in-tensity of illumination. In this method, the sum ofthe rates of photochemical reactions I and II is mea-sured by the difference in the rate of membrane po-tential changes determined immediately before or afterswitching off the light. Experiments were performedunder strong light excitation in the presence of air, withdark-adapted spinach (Joliot and Joliot, 2002) or Ara-bidopsis 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 illu-mination, rate of the cyclic flow is estimated to ∼130sec−1 and remains roughly constant during the first 10sec of illumination. Unexpectedly, this cyclic processis not inhibited by antimycin (Joliot and Joliot, 2002).Under the same conditions, fluorescence induction ki-netics shows that a pool of soluble PS I acceptors (Fd,FNR and NADP) of approximately nine electron equiv-alents is reduced in less than 100 msec via the linearpathway. This implies that, even in dark-adapted leaves,PS I remains able to transfer electrons to a pool of oxi-dized PS I acceptors. In the presence of DCMU, wherethe linear flow is fully inhibited, a similar rate of thecyclic flow is measured during the first seconds of illu-mination. This rate progressively drops to zero in ∼7sec, due to slow electron leaks that lead to the oxidationof the carriers involved in the cyclic chain.

After 100 msec of illumination at an intensity thatis saturating for the cyclic process, most of P700 isreduced (Harbinson and Hedley, 1993; Strasser et al.,2001; Joliot and Joliot, 2002; Schankser et al., 2003)implying that a fast charge recombination betweenP700+ and reduced acceptors will occur. In agreementwith this assumption, it is observed that the kinetics ofthe membrane potential displays a fast decaying phaseof small amplitude, which is completed in ∼500 μsec.The amplitude of this phase is roughly proportionalto the light intensity. This phase correlates with a re-duction phase of P700+ (measured by the absorptionchanges at 810 nm) and has been ascribed to a chargerecombination between P700+ and most probably theiron-sulfur carrier FX (Joliot and Joliot, 2002). It canbe thus concluded that most of the carriers of the linearand cyclic chains are poised in their reduced state. Asmall fraction of “cyclic PS I” centers includes P700+

and a single negative charge on the FA/FB cluster. Forthese RCs, the rate of P700+ reduction via the cyclicpathway is faster than the rate of charge recombinationbetween P700+ and (FA/FB)− (t1/2 ∼ 45 msec; Hiyamaand Ke, 1971).

A charge recombination process is also observed inthe presence of DCMU, the amplitude of which is half


that measured in its absence. This is explained by thefact that illumination induces the oxidation of all thecarriers of the linear chain, while the carriers involvedin the cyclic pathway are transitorily poised at a poten-tial able to induce the reduction of most of the FA/FB

acceptors (<−600 mV). In the 1–7 sec time range, thedecrease in the rate of the cyclic flow is associated witha corresponding decrease in the amplitude of the chargerecombination phase. This parallel decrease reveals aprogressive oxidation of all the carriers involved in thecyclic process, associated with a slow electron leak to-ward O2 or the Benson–Calvin cycle.

VII. Pathway of Cyclic Flow in HigherPlants

Work described above leads us to conclude that cyclicET is a real physiological phenomenon, though per-haps only occurring under a limited range of conditions.The question then arises, what is (are) the physiologi-cal pathway(s) for this electron flow. Early work indi-cating two pathways of cyclic ET in isolated systemswere mainly based on measurements of chloroplastsisolated from C3 plants. We would therefore expect tosee evidence for both the ferredoxin-linked antimycin-sensitive and the NADPH-linked antimycin-insensitivepathways in whole leaves. The in vivo data on theeffect of antimycin A are however, ambiguous. Joetet al. (2002) reported a slow cyclic ET measured underfar-red excitation is stimulated in anaerobiosis but alsoin the presence of inhibitors of the respiratory chain,including antimycin. In the same way, antimycin doesnot inhibit the fast cyclic flow measured under strongillumination of dark-adapted leaves (Joliot and Joliot,2002). In both cases, it was proposed that the effectof antimycin is related to the inhibition of the respi-ratory chain, which increases the reducing power inchloroplasts via mitochondria–chloroplast interac-tions. Further confusion might arise in in vivo mea-surements because of other functions of antimycin, no-tably its ability to inhibit nonphotochemical quenching(Oxborough and Horton, 1987). We thus conclude thateffect of antimycin is not a decisive test in proposingor excluding the occurrence of cyclic electron flow.

A. NADPH-Dependent Cyclic ElectronTransfer

The more convincing arguments that favor the involve-ment of NADPH:PQ reductase (NDH) in the cyclicpathway come from the analysis of mutants lackingthis enzyme. NDH-dependent cyclic ET reported in the

literature appears as a slow process that is very likely re-lated to the low concentration of NDH [∼1% of that ofthe photosynthetic chain (Sazanov et al., 1995)]. Levelsof this complex are seen to increase under conditionsof stress, suggesting a role in survival under conditionsof oxidative stress (Casano et al., 2000; Teicher et al.,2000; Lascano et al., 2003), however, to what extent thisincreases the flux through the NDH complex remainsunclear. Burrows et al. (1998) observed that plantslacking the NDH complex lacked the transient fluo-rescence rise seen following illumination. Hashimotoet al. (2003) were recently able to use this character-istic to identify a new mutant of A. thaliana, CRR2,which lacks a nuclear encoded factor that seems to beessential for NDH expression. However, the absenceof a fluorescence rise in NDH-less mutants seems tobe highly sensitive to the developmental and metabolicstatus of the leaf (A. Krieger-Liszkay, personal com-munication.), supporting the idea that this is not theonly route for reduction of the PQ pool by stromalfactors. Evidence has also been published that the re-duction of P700+ following far-red light is affected inplants lacking the NDH complex (Joet et al., 2002).Involvement of NDH in the fast cyclic ET measured indark-adapted leaves submitted to strong illumination(Joliot and Joliot, 2002, 2004) appears unlikely as thisenzyme, which is present at low concentration, wouldhave to operate at rate as high as ∼104 sec−1 to sustaina rate of electron flow of 130 sec−1.

B. Ferredoxin-Dependent Cyclic ElectronTransfer

If rapid rates of cyclic ET cannot be sustained by theNDH complex, then we must consider the likely in-volvement of a ferredoxin pathway. Electrons maybedirectly 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 mechanismis proposed that results in the pumping of one protonper electron transferred, but differs from conventionalQ-cycle process (Joliot and Joliot, 2004). FNR isknown to copurify with the cyt b/ f complex (Clarket al., 1984) and provides a binding site for Fd (Zhanget al., 2001). It seems likely that the electron path-way between Fd and site Qi involves the covalentlybound cyt c′ (cyt ci) recently characterized in the struc-ture of cyt b/ f complex in Chlamydomonas reinhardtii(Stroebel et al., 2003) and in cyanobacteria (Kurisuet al., 2003). On the other hand, it has been proposedthat an electron carrier G (Lavergne, 1983; Joliot andJoliot, 1988), is localized on the stromal side of the cytb/ f complex and able to exchange electrons with cyt


Fig. 2. A mechanism for the ferredoxin-dependent electrontransfer.

bh. This carrier G has been now identified as cyt ci

(Alric et al., 2005). According to Fig. 2, sites Qi andQo behave quite symmetrically. At site Qi, reductionof PQ involves the transfer of one electron from Fdand one electron from cyt bh; at site Qo, oxidation ofPQH2 involves the transfer of one electron to cyt f viathe Rieske Fe/S protein and one electron to cyt bl. Itworth noting that, in the dark, reduction of the PQ poolcould be induced by the sequential transfer of electronsfrom Fd to site Qi and the cyt b/ f complex would thenbehave as a FQR. Evidence for a distinct FQR, indepen-dent of the Qi site, is related to the effect of antimycincompared to HQNO (Moss and Bendall, 1984). Thisevidence only rules out a role for antimycin in blockingthe Qi site and does not rule out the involvement of thatsite, if it does not bind antimycin.

It has been proposed that the small hydrophilicpolypeptide PGR5 is required in a process of protongradient generation and could be involved in FQR ac-tivity (Munekage et al., 2002). At present, very littleinformation on this mutant is available and a better

characterization of its function is required. PGR5 lacksa metal binding motif that might implicate it in a directrole in electron transfer, however it might play a sec-ondary role, e.g., in being involved in the binding ofFNR to the cyt b/ f complex. The published evidencethat PGR5 is involved in cyclic ET is based on the obser-vation that NADPH and Fd-dependent reduction of PQpool, measured by chlorophyll fluorescence is impairedin PGR5 mutant. This observation needs to be consid-ered with care, however, since the effect on the kineticsof PQ reduction and the sensitivity to antimycin appearto be qualitatively the same, with only the amplitude ofthe fluorescence rise being changed (Munekage et al.,2002). Since the latter might be sensitive to other fac-tors, for example the presence of photoinhibited PS IIcenters in the thylakoid membrane, this result is notconclusive.

The fraction of PS I centers that operate accordingto the cyclic or linear mode has been determined bymeasuring P700 and PC oxidation under weak far-redexcitation (8 photons / PSI / s), a photochemical rateconstant well below the rate constant of the limitingsteps of cyclic or linear flow (Joliot and Joliot, 2005).Far-red illumination of a dark-adapted leaf induces aslow oxidation of P700 and PC that is completed in10–20 sec. During this oxidation phase, the number ofPS I turnovers is much larger than the number of elec-trons stored in the primary and secondary PS I donors.It implies that most of the electrons formed on the stro-mal side of PS I are transferred back to PS I via thecyt b/ f complex (cyclic electron flow). Slow electronleaks, probably toward the Benson-Calvin cycle, resultin a progressive oxidation of most of the carriers in-volved in the cyclic pathway, including P700. When thesame leaf is first preilluminated for several minutes un-der green light that excites both photosystems and acti-vates the Benson-Calvin cycle, P700 oxidation inducedby far-red excitation is a much faster process completedin less than 3 sec. The kinetics of P700 oxidation mea-sured with a preilluminated leaf is close to that mea-sured in the presence of methyl viologen, an efficientPS I electron acceptor. It implies that, in preillumi-nated leaves, most of the electrons reaching the stromalsize of PS I are transferred to the Benson-Calvin cy-cle via FNR and NADP (linear electron flow). Usingthe same approach, the transition from a cyclic (dark-adapted leaf) to a linear mode (preilluminated leaf) hasbeen recently analyzed during the course of the greenpreillumination (P. Joliot, unpublished data). For thefirst minute of illumination, most of PS I centers con-tribute to cyclic electron flow. During the subsequentperiod of illumination (1–10 min), activation of the


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 therate of linear flow, correlates with a decrease in the rateof the cyclic flow. In the same way, the slowdown ofthe linear flow induced by a lack of CO2 is associatedwith an increase of the rate of the cyclic flow, as previ-ously proposed by Heber and co-workers (Hauser et al.,1995). These results suggest that linear and cyclic path-ways be in permanent competition for the reoxidationof Fd. The relative efficiency of cyclic and linear path-ways is suggested to be determined by the probability ofFd binding either to a site localized on the stromal siteof the cyt b/ f complex or to an ‘active’ FNR. This wedefine as an FNR molecule to which NADP+ is bound.Activation of cyclic flow is thus associated with the re-duction of NADP+ and the competition between cyclicand linear mode will be controlled by the redox stateof NADP which depends upon the degree of activationof the Benson-Calvin cycle. In dark-adapted leaves, il-lumination first induces the reduction of the small poolof acceptors present in their oxidized state (FNR andNADP+). When this pool is fully reduced, Fd in excessis available to bind the stromal site of cyt b/ f , whichinitiates cyclic electron flow.

In the model depicted in Fig. 3, Fd is presumed tofreely diffuse in the stromal compartment. This con-trasts with models in which the cyclic chain is orga-nized in supercomplexes that associate 1 PS I center / acyt b/ f 1 PC / 1 Fd (Carillo and Vallejos, 1983; Laisket al., 1992; Laisk, 1993; Joliot and Joliot, 2002). Inthis hypothesis, the concentration of PS I included inthe supercomplex cannot be larger than that of cyt b/ fcomplex localized in the non-appressed region of themembrane. 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 re-gion Albertsson, 2001), the obligatory formation of su-percomplexes for cyclic electron flow can be excluded.

VIII. Cyclic Flow in Green UnicellularAlgae

There is a general consensus that a cyclic electron flowoperates in algae, at least under anaerobic conditions,when PS II activity is mainly inhibited due to the re-duction of QA. It worth pointing out that, in contrast tohigher plants, anaerobiosis is a common occurrence formany algae in their natural environment. Anaerobiosisis known to induce state 1 to state 2 transition associ-ated with changes in the distribution of the membranecomplexes (Bulte et al., 1990) to a much larger extentthan those induced by chromatic adaptation (Bonaven-tura and Myers, 1969). Anaerobiosis, like all treatmentsthat decrease the ATP content of the cell (mitochon-drial inhibitors, uncouplers, ATP-synthase inhibitors),induces the transfer of most of the light harvesting com-plex LHC II (Bulte et al., 1990) and of a fraction ofcyt b/ f complexes (Vallon et al., 1991) from the ap-pressed to the nonappressed region of the thylakoids.On the basis of measurements of the turnover rate ofcyt f , Finazzi et al. (1999) concluded that only linearET is active in the presence of O2 (state 1 conditions),while cyclic electron flow operates in state 2 condi-tions (anaerobiosis or presence of uncouplers), wherethe linear process is fully inhibited. The rate of cyclicflow measured in state 2 conditions is similar to that


measured for the linear flow at the same light inten-sity in state 1 conditions. One can thus conclude that inanaerobic conditions, most of the antennae is involvedin the cyclic process with ∼80% of the light collectedby the “cyclic PS I.” In these conditions, unicellularalgae may behave in a similar way to the green bac-teria with type I photosystems, with a photosyntheticprocess entirely devoted to ATP synthesis. The cyclicprocess in algae thus appears fundamentally differentfrom that in higher plants, which occurs in the presenceof O2 and state 1 conditions.

A. Mechanisms of Cyclic Flow in Algae

Complexes containing PS I and cyt b/ f complexeshave been observed in solubilized membranes ofC. reinhardtii (Wollman and Bulte, 1989) possiblypointing to the presence of supercomplexes in this or-ganism. On the other hand, the analysis of ET kineticsin state 2 conditions in a mutant with a low cyt b/ f con-tent has shown that PC is able to freely diffuse from anyPS I center to cyt f (Finazzi et al., 2002). This excludesthe presence of functional supercomplexes, includinga trapped PC. In state 2 conditions, a spatial separa-tion between the linear and cyclic chains results fromthe transfer of a large fraction of the stromal cyt b/ fcomplexes from the appressed to the nonappressed re-gion. Thus, one expects that, even in the absence ofsupercomplexes, an efficient cyclic ET will occur inthe nonappressed region that includes the PS I centersand most of the cyt b/ f complexes.

The electron pathway from PS I to the cyt b/ f com-plex has not been identified. While no gene encodingfor an enzyme similar to NDH has been identified inthe chloroplast of algae, the presence of a PQ reduc-tase is suggested by the slow reduction of PQ poolin the appressed region, which has been observed inthe presence of O2 (Diner and Mauzerall, 1973). Oneexpects that such an enzyme would operate at a muchhigher rate in anaerobic conditions. This enzyme couldbe involved in a cyclic process if it were present in thenonappressed region. Another possible pathway is a di-rect ET from PS I to cyt b/ f complex, as depicted inFig. 1 (pathway 3). It worth pointing out that, to ourknowledge, complexes that associate FNR and the cytb/ f complex have not been identified in C. reinhardtii.

IX. Cyclic Flow in Cyanobacteria

In cyanobacteria, the presence of respiratory and pho-tosynthetic electron transport chains in the same mem-

brane system, to the point where the two share redoxcomponents (Scherer, 1990), means that the cyclic EToccurs in an environment very different to that in plantsand algae. A common feature to most of cyanobacte-riae is the large excess of PS I as compared to PS IIcenters (Fujita et al., 1994). This suggests that evenunder steady-state condition of illumination, a frac-tion of PS I centers could be involved in cyclic ET.On the other hand, in cyanobacteria, PS I centers areorganized in trimers, which exclude the formation ofsupercomplexes that associate PS I centers with the cytb/ f complex.

After illumination in the presence of DCMU or af-ter far-red excitation, reduction of P700+ occurs witht1/2 ∼ 500 msec, taken to be indicative of cyclic ET(Maxwell and Biggins, 1976; Mi et al., 1992a,b; Yu etal., 1993). At the same time, measurements of energystorage via photoacoustics point to the occurrence ofcyclic ET under such conditions. However, it can be as-sumed that, in dark-adapted samples, the PQ pool willbe maintained in a relatively reduced state due to respi-ratory electron flow. It needs to be shown, therefore, thatelectron transport from the acceptor side of PS I backinto the PQ pool occurs and is maintained under con-ditions where PS II is active. (Sandmann and Malkin,1983, 1984) were able to observe that, while respira-tion was inhibited in the light, due to competition withphotosynthesis, NAD(P)H oxidation by spheroplastsfrom Aphanocapsa was less affected. The addition ofDCMU did not enhance NAD(P)H oxidation, suggest-ing no competition between NDH and PS II for PQreduction. However, as pointed out by Mi et al. (2000),the organization of the membrane systems in cyanobac-teria may be substantially disturbed in isolated systems.As far as we are aware, these results have not yet beenconfirmed in intact cells.

Measurements on mutants lacking the NDH com-plex suggest this is generally the primary route forcyclic ET (Yu et al., 1993; Mi et al., 2000). Mutantslacking ndhF of the NDH complex and psaE in PSI, show inhibited growth at low-light intensities, butare unaffected at high light. Cyclic ET in cyanobacte-ria has been implicated in providing energy for CO2

concentrating mechanisms. Growth of Synechocystisat low CO2 results in upregulation of NDH subunits(Deng et al., 2003). Mutants lacking the NDH com-plex have a reduced ability to tolerate growth underlimiting CO2 conditions (Ogawa, 1991). On the otherhand, it has been reported that Synechocystis sp. PCC6803 contains a succinate quinol oxidoreductase that issuggested to contribute to cyclic electron flow (Cooleyet al., 2000). This enzymatic activity would, however,


involve a more complex cyclic pathway than so farconsidered and it is questionable whether it can still beregarded as cyclic electron flow.

Exposure to high salt concentrations leads to an ac-tivation of a cyclic ET pathway that does not requireNDH (Jeanjean et al., 1998). The details of this pathwayremain unclear but probably involve FNR (van Thoret al., 2000; Matthijs et al., 2002). Binding of the FNRto the membrane is facilitated by an N-terminal domainwhich is absent in the higher plant enzyme (van Thoret al., 2000). Low concentrations of sodium bisulfitehave also recently been suggested to stimulate cyclicET (Wang et al., 2003).

X. Functions and Regulation of CyclicElectron Transfer

The primary role of cyclic ET has always been supposedto be the generation of ATP, either to support CO2 fixa-tion or for other metabolic processes. The requirementfor cyclic ET to balance the production of reductantand ATP in normal CO2 fixation conditions has beenwidely debated. Structural analysis of the electrical ro-tor of the chloroplast ATP-synthase has shown that it ismade up of 14 polypeptides (Seelert et al., 2000). Onethus expects that a complete rotation of the rotor, whichinduces the synthesis of 3 ATP, is associated with thetransfer of 14 protons across the membrane that leadsto H+/ATP ratio of 14/3 = 4.7. Assuming that 3 protonsare 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 asion leaks through the membrane are not taken into ac-count. This amount of ATP is definitely lower than thatrequired by the Benson–Calvin cycle (3 ATP/CO2). Itis thus likely that, even in light-adapted conditions, asmall fraction of PS I contributes to a cyclic flow tosatisfy the ATP requirement of the Benson–Calvin cy-cle. This could explain why the minimum quantum re-quirement of the photosynthetic process measured un-der weak excitation (10–12 quanta/O2) is significantlylarger than the theoretical value of eight quanta/O2.

If cyclic ET is contributing ATP to drive CO2 fixationand assuming that this is regulated in a way that ensuresthat the ATP supply is maintained at a constant level,then we would expect that under steady-state conditionof illumination, the rate of cyclic ET would parallel therate of linear ET. Thus, it is perhaps unsurprising thatlight-limited steady-state measurements, which rely oncomparing PS I and PS II ET, have typically found noevidence for this cyclic flow (Harbinson et al., 1990;

Sacksteder and Kramer, 2000). By contrast, when theATP consumption is not directly coupled to CO2 fix-ation, we would expect to find discrepancies betweenPS I and PS II turnover. This would certainly be thecase in C4 bundle sheath cells, where the reductant re-quired for CO2 fixation is not provided by linear ET. Itwould also be the case during the first few seconds ofillumination, during which time a trans-thylakoid pHgradient must be established before ATP synthesis canbegin (Joliot and Joliot, 2002).

In green algae, reducing conditions (anaerobiosis)result in a large state transition, with phosphorylationof LHC II resulting in its migration to PS I, push-ing the chloroplast into a “cyclic-only” state. Underanaerobiosis, mitochondrial respiration will be inhib-ited, leading to a deficit of ATP and possibly also to arise in the concentration of NADH. A shift from linearto cyclic photosynthetic ET would counteract this ef-fect, lowering the production of reducing potential infavor of ATP synthesis. Thus the redox regulation ofstate transitions in algae and the resultant regulationof cyclic ET is a mechanism for balancing the overallcellular ATP/NAD(P)H ratio.

In higher plants, state transitions are less prominentand their regulation in response to redox potential verydifferent. Reducing conditions induced by anaerobio-sis do not give rise to state 1 to state 2 transitions butthey 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 con-trast to algae, efficient cyclic ET operates during thefirst seconds of illumination of dark-adapted leaves (Jo-liot and Joliot, 2002), i.e., in pure state 1 conditions,and there is no evidence that state transitions lead toan increase in cyclic ET, rather they are thought to bal-ance excitation of PS I and PS II for optimal linearET. The small state transitions that do occur are seenat very low light and depend on the redox state of thePQ pool. On the other hand, the observation that A.thaliana grown under very low-light increase PS I rel-ative to PS II might suggest a role for cyclic ET undersuch conditions, implying a switch from CO2 fixationto ATP synthesis (Bailey et al., 2001).

A notable difference between higher plants, somealgae and cyanobacteria is in their ability to protectthemselves from high light through the process of highenergy state quenching (qE). This process, which islinked to the presence of the carotenoid zeaxanthin,is driven by �pH. It was first suggested some yearsago that the �pH required to generate this quenchingwas generated by cyclic ET (Heber and Walker, 1992);however, it is only recently that direct evidence for its


requirement has been obtained. The PGR5 mutant of A.thaliana that is thought to be deficient in cyclic ET, wasisolated using a screen that selected for a deficiencyin quenching (Munekage et al., 2002). Golding andJohnson observed a quantitative link between the extentof non-photochemical quenching (NPQ) and the extentof 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 pathwaysof cyclic ET may be fulfilling different functions andtherefore be differentially regulated. Upregulation ofcyclic at low CO2 correlates with a downregulation ofthe cyt b/ f complex, inhibiting linear ET (Golding andJohnson, 2003). This downregulation of linear ET hasbeen suggested to be regulated via thioredoxin (John-son, 2003). Thus, upregulation of cyclic ET in responseto reducing conditions seems likely, although the mech-anism involved is different to that seen in green algae.

Joliot and Joliot (2002) suggested that the cyclic ETseen following a dark to light transition in dark-adaptedleaves maybe regulated by the ATP/ADP ratio in thechloroplast. This is a reasonable suggestion, if we ac-cept that this cyclic is required to kick-start ATP syn-thesis. On the other hand, the cyclic ET seen under suchconditions has a short lag period, during which time theFd and NADPH pools are likely to become reduced,in agreement with a redox regulation process. After apreillumination, the transition from the linear to thecyclic mode requires more than 1 hour of dark adap-tation (Golding et al., 2004; Joliot and Joliot, 2005).Deactivation of the Benson-Calvin cyclic following il-lumination is slow, taking up to 3 hours or more in somespecies. Thus the long period of time required for theinduction of a cyclic activated state might be explainedsimply by this deactivation, though changes in the en-ergy status of the cell, related to the consumption ofcarbohydrate reserves might also play a role.

Given the low requirement for additional �pH aris-ing from CO2 fixation, it might be speculated that thecapacity of NDH pathway is sufficient to support this.However, mutants lacking NDH are clearly not defi-cient in CO2 fixation. It may be however that a nor-mally stress (redox) activated cyclic pathway becomespartially activated under conditions where NDH is lack-ing, compensating for the lack of this pathway. Thereis little evidence that the NDH pathway contributes tophotoprotection. Ndh-less mutants have normal lev-els of NPQ and are not more susceptible to photoin-hibition either of PS I or PS II at ambient or lowtemperature (Barth and Krause, 2002). On the otherhand, there are some indications that plants lacking

NDH are more sensitive to water stress (Horvath et al.,2000).

In cyanobacteria, there is a large amount of data thatlink the NDH complex to CO2 concentrating mech-anisms. It seems that there are distinct forms of thecomplex that are involved in this process, with therepossibly being a direct role of NDH in the formation ofHCO−

3 from CO2. It is not clear how this relates cyclicET however, i.e., whether this reaction depends on elec-tron transport occurring through PS I (see discussionin Badger and Price, 2003).

XI. Conclusion

The pathway of linear electron transport has beenclearly established for many years. Cyclic ET on theother hand remains enigmatic. Nevertheless, this is anarea in which significant progress is beginning to bemade. The establishment of new methods for the quan-titation of cyclic flow, such as those outlined here, willhelp in this progress. Clearer ideas on the function ofcyclic ET will also help in the identification of the pro-teins involved in this pathway, either through conven-tional screening approaches or through reverse genet-ics, examining the effects of knockout mutations oncyclic electron transport.

Acknowledgments

We would like to thank Dr. Giovanni Finazzi (IBPC,Paris, France) and Dr. Conrad Mullineaux (Univer-sity College, London, UK) for useful discussions andProf. Toshiharu Shikanai (Nara Institute of Science andTechnology, Japan) for providing a reprint of his work.

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