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Photosynthesis and chlororespiration - competition orsynergy ?

Wojciech Jacek Nawrocki

To cite this version:Wojciech Jacek Nawrocki. Photosynthesis and chlororespiration - competition or synergy ?. VegetalBiology. Université Pierre et Marie Curie - Paris VI, 2016. English. �NNT : 2016PA066599�. �tel-01599264�

PhD Thesis of the Pierre and Marie Curie University (UPMC)

Prepared in the Laboratory of Molecular and Membrane Physiology of the Chloroplast, UMR7141, CNRS/UPMC

Doctoral school: Life Science Complexity, ED515

Presented by Wojciech Jacek Nawrocki for the grade of Doctor of the Pierre and Marie Curie University

Photosynthesis and chlororespiration

– competition or synergy?

Defended on the 30th of September, 2016 at the Institute of Physico-Chemical Biology in Paris, France

PhD Jury:

Anja Krieger-Liszkay, CEA Saclay, reviewer

Pierre Cardol, University of Liège, reviewer

Roberto Bassi, University of Verona, examinator

Giles Johnson, University of Manchester, examinator

Sophie Cribier, UPMC, examinator

Francis-André Wollman, CNRS/UPMC, PhD supervisor

Pierre Joliot, CNRS/UPMC, invited member

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Dla Mamy i Taty

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Acknowledgements

Thanks to the late Fabrice Rappaport, my PhD supervisor. He is truly missed and will

be looked up to forever.

Thanks to Francis-André Wollman, for having me in the lab since that Erasmus

internship until today – but before all, for the discussions and support. I am honored

to have been a part of your laboratory.

Thanks to Pierre Joliot and Benjamin Bailleul – I would know infinitely less without

you; and it was invariably a pleasure to talk.

Thanks to my colleague and friend, Stefano Santabarbara.

Thanks to Stephan Eberhard, for a great time during my M1 internship and for all the

advice.

Thanks to my collaborators, Felix Buchert, Nicolas Tourasse, Laura Mosebach,

Antoine Taly and, in particular, Pierre Sétif. It was a privilege to work with you.

Thanks to all other people from the lab for making it such a great place.

I am indebted to the honorable members of the jury – thanks for accepting this task.

Thank you, Wojtek 2, for all your help and for the fun. It was never boring.

Thanks to Larissa for your love and support. ♡

Finally, thanks to my family for all kinds of help.

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Table of Contents

Introduction to photosynthesis ............................................................................... 9

Opening words ...................................................................................................................... 9 Primary processes ................................................................................................................13 Regulation of photosynthesis ................................................................................................22

PTOX and chlororespiration .................................................................................. 31

Historical view ......................................................................................................................31 PTOX review ........................................................................................................................37 New data about PTOX in the literature .................................................................................81

Results and discussion .......................................................................................... 84

Chlororespiration and its physiological role ...........................................................................84 Physiological observation .................................................................................................84 PTOX in fluctuating light paper .........................................................................................88

Trials of quantitative in vivo measurements of chlororespiration reveal information about CEF in Chlamydomonas ............................................................................................................. 120

Initial idea overview – measurements of chlororespiratory fluxes .................................... 120 Results ........................................................................................................................... 130 Implications for CEF mechanism .................................................................................... 151

State transitions .................................................................................................................. 176 Introduction ..................................................................................................................... 176 State transitions paper .................................................................................................... 179 Perspectives, analysis of discrepanicies between labs ................................................... 217

Conclusions and perspectives ............................................................................ 222

References ............................................................................................................ 231

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Introduction to photosynthesis

Opening words

Photosynthesis sustains the vast majority of life on the Earth. This process consists

of the light absorption by plants, algae and cyanobacteria and conversion of the

kinetic energy of photons (with a wavelength dependence given by the Planck-

Einstein relation 𝐸 = ℎ𝑣) into highly energetic bonds in organic molecules. It allows in

parallel the reduction of carbon (IV) from CO2 into lower oxidation states, which are

useful for respiration. Finally, nearly as a side reaction, ATP – the universal energy

currency of the cell - is synthesized in the process.

The first discoveries concerning the topic can be dated back to the 17th century when

Van Helmont followed growth of plants only provided with water. Since then, we know

infinitely more about this process. A little part of the most crucial information

concerning photosynthesis will be presented here.

Photosynthesis first appeared in what we now call purple bacteria, a class of

prokaryotes inhabiting extreme environmental niches. The early types of this process

were anoxygenic, i.e. they did not result in the production of O2 as a byproduct. The

ability of using water as an electron donor evolved first in cyanobacteria around 3,5

By ago (fig. 1).

Since then, the ability to perform oxygenic photosynthesis greatly spread. A vast part

of the photosynthetic activity on Earth takes place in chloroplasts, intracellular

organelles with a double membrane envelope. As first proposed by Mereschkowski in

1905, chloroplasts are ancestors of a cyanobacterium that was engulfed by a

eukaryotic cell. The to-be organelle lost a massive part of its genome, with a lot of

genes being transferred to the nucleus of the host as depicted in the fig. 1.

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Fig. 1 The evolution of plastids and mitochondira (Adapted from (Dyall et al, 2004))

Because of this horizontal gene transfer from the endosymbiont to the nucleus, the

photosynthetic machinery is now partly coded by the nucleus, and partly by the

chloroplast genome. Because of differences in the two genome poliploïdy, the

establishment of the proper ratio of subunits required to build up each functional

photosynthetic oligomeric complexes, is attained through the tight control of the

chloroplast expression machinery by a set of trans-acting protein factors encoded by

the nucleus.

Photosynthesis is a process tightly associated with membranes. The chloroplasts,

although their structure is not identical with regards to cyanobacterial cells, share a

similarity with the latter. Both are filled with scaffolds of membranes where

photosynthesis takes place. In the case of chloroplasts of cyanobacteria, algae and

plants, those structures are called thylakoids and consist of lamellae membranes,

which are unstacked, and grana stacks in which the membranes are tightly packed.

This structuration is shown in fig. 2. The interior of membrane vesicles is called

lumen, while the bulk of the chloroplast is its stroma.

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Fig. 2 Structure of the chloroplast. Top, a comparison between chloroplasts and

cyanobacteria. Left, Transmission electron microscopy images of Chlamydomonas

chloroplasts where stacks of membranes are visible. gt, grana thylakoids. Right, 3D

reconstruction of the thylakoid structure deduced from the electron microscopy. Light

grey, grana stacks. Colored, stroma lamellae. Adapted from: Wikipedia, CC,

Kelvinsong; (Daum et al, 2010); (Daum & Kuhlbrandt, 2011)

The chemistry of oxygenic photosynthesis is well established. The overall reaction

catalyzed can be described as follows:

CO2 + H2O + photons → [CH2O] + O2

Hence, photosynthesis transfers electrons from oxygen (-II) to carbon in CO2 (IV).

This reduction of atmospheric CO2 allows plants to use it as a carbon source (during

growth), as well as the energy source (to sustain metabolic processes).

Photosynthesis therefore catalyzes a reaction which is unfavorable energetically, i.e.

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has a positive Gibbs free energy change of ∆G°’ = 2867kJ/mol. The energy, which is

used to oxidize a strongly electronegative atom - the oxygen from water molecules -

comes from light. In order to allow spontaneous transfer of electrons toward the

oxidized carbon, the former are excited by photons. The excited pigments have

strongly negative redox potentials, i.e. have the tendency to donate electrons. Light-

driven reactions in photosynthesis are able to yield compounds with potentials

reaching -1200 mV, allowing the subsequent reduction of NADP+ to NADPH (-370

mV in physiological conditions) and making the reduction of CO2 possible.

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Primary processes

Light absorption

Photosynthetic cells developed specialized structures to take advantage of the

largest quantity of light available. The chloroplasts, in particular their thylakoid

membranes, are dark green in colour because of the quantity of pigments embedded

inside the bilayers. The most important pigment is chlorophyll, however carotenoids

are also abundantly present in the membrane.

Photosynthetic pigments are held in protein scaffolds in the membranes. We can

distinguish two types of pigment-binding proteins – antenna complexes (also known

as light-harvesting complexes, LHC), the role of which is essentially to capture light

energy, and transfer it to the second type of pigment-protein complexes, the

photosystems. The latter are complex photochemical enzymes, which will be

described below.

By the mean of absorption, antenna pigments transform the kinetic energy of photons

into their internal energy. This is because upon photon absorption an electron of the

pigment is moved to a different shell (i.e. one of its quantum numbers changes from

the minimal ground state)(Fig. 3). The pigments in the relatively rigid antennae are

held in place in distances allowing a fast transfer of the excitation energy, which can

be described either with Förster or Redfield theories (Fig. 3, III)(Croce & van

Amerongen, 2014).

The excitation energy is funnelled to the reaction centre of photosystems in sub-

nanosecond scale, however if it is not rapid enough the excitation can decay (Fig. 3, I

and II).

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Fig. 3. Possible scenarios after a molecule absorbs light energy.

Charge separation

Photosystems are large, multimeric complexes, which drive charge separation. When

the exciton reaches the so-called special pair of chlorophylls, i.e. its reaction centre,

the excitation transfer no longer occurs, but the electron in excited state is transferred

from the chlorophyll to its acceptor, i.e. trapped. Thanks to proximity and positioning

of the acceptors and their consequently increasing redox potentials inside the low

dielectric constant photosystem scaffold, the “forward” transfer is preferential with

regards to a decay of the exciton or a recombination of the charges (Fig. 3, IV).

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Fig. 4. Structure of the active centers and cofactors involved in charge separation

and stabilisation of Q-type (quinone) and Fe-S type photosystems. Q-type reaction

center of Cyanobacteria is the same as in photosynthetic eukaryotes. From

(Hohmann-Marriott & Blankenship, 2011)

There are two types of photosystems in oxygenic photosynthesis (fig. 4). Both

operate with similar principles yet use variants of terminal electron acceptors.

Photosystem II (PSII) is located for its vast majority in the appressed regions of the

membrane, the thylakoid grana. It uses quinones, organic molecules with a

hydrophobic chain soluble in the membrane, as electron acceptors and can be

described as a water:plastoquinone oxidoreductase. The complex physical chemistry

of the oxidation of water, which results in a release of protons and oxygen to the

lumen and a reduction of the oxidized special pair, is a unique feat in the biological

world and is a subject of an ongoing debate over its precise mechanism. PSII forms

supercomplexes with its antenna, LHCII.

Photosystem I (PSI), on the other hand, uses iron-sulfur clusters as terminal

acceptors, and is a plastocyanin:ferredoxin oxidoreductase mostly present in the

stroma lamellae. Antennae of PSI are called LHCa. Reduced Ferredoxin produced by

the PSI in turn provides electrons used by the enzyme Ferredoxin:NADP+

oxidoreductase to form NADPH.

Electron transfer and the Q-cycle

PSII transfers sequentially two electrons to a plastoquinone molecule. Upon its

reduction, taking place at the QB binding site of PSII, the two additional electrons are

compensated by a protonation. Protons for this reaction are taken up from the stroma

of the chloroplast.

The newly created non-polar dihydroxy-plastoquinone, or plastoquinol (PQH2) then

diffuses in the tightly packed thylakoid membrane to the plastoquinol:plastocyanin

oxidoreductase, cytochrome b6f. This complex enzyme is responsible for the so-

called Q-cycle, a mechanism that essentially allows conservation of potential energy

– which is released by electrons reducing compounds with increasing redox potential

(as depicted on the Z-scheme of photosynthesis, fig.5):

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Fig. 5. The Z-scheme of photosynthesis. This diagram depicts the cofactors involved

in transfer of electrons from water to NADPH and links each of them to the midpoint

redox potential of its oxidized/reduced couple. The y-axis is inversed in order to allow

a ‘gravitational’ analogy in which the electrons spontaneously ‘fall’ to cofactors with

more positive potential, whereas light energy is needed to ‘lift’ the elctrons, i.e.

produce an unstable, reducing excited state with relatively negative potential.

Upon reaching the quinol oxidation site in b6f (QO)(fig. 6), the two-electron transporter

first reduces, in an uphill reaction, the iron-sulfur cluster of the Rieske protein, which

in turn reduces cyt. f and then the acceptor, plastocyanin (photooxidized by the PSI).

This reaction releases two protons to the lumen. Quinone oxidation is a limiting

reaction in the electron transfer when the light is abundant, limiting the flow to 150 e-

/s/photosystem. The semiquinone produced by the oxidation has a low redox

potential and can reduce the low potential chain of the b6f consisting of three haems

– conserved in b6f’s mitochondrial counterpart bc1 haems bl close to luminal, and bh

close to the stromal side of the membrane (see fig. 6). An additional ci haem,

positioned next to the bh is found exclusively in cyt. b6f (Stroebel et al, 2003). After

this reaction repeats, those two haems have accumulated one electron each and can

in turn reduce a quinone molecule at the Qi site, which becomes electrically neutral

upon a double protonation by H+ ions from the stroma. Plastocyanin in the next step

of electron transfer reduces the oxidized special pair of the PSI.

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Thanks to this design, initially proposed by P. Mitchell, a part of the electrons’ energy

is used to translocate protons from the stroma to the lumen.

Fig. 6. Cytochrome b6f operates through Mitchell’s modified Q-cycle. Left side, a

schematical representation of the Q-cycle. The b6f complex transfers two protons

(green arrows) per one electron transferred (blue arrows) along high (Fe2S2 cluster,

cytochrome f) and low potential chains (bl, bh, ci haems). Quinol (QH2) oxidation at Qo

site, Quinone (Q) reduction at Qi site. Golden arrows, quinone movements; according

to Joliot & Joliot (xx), the quinone formed at the Qo site has direct access to the Qi

site without an exchange with the PQ pool. Top right side, vicinity of the stromal

haems (Qi site) seen from the top of the complex. Right side, structure of a b6f

monomer and its cofactors: purple, b haems; red, ci (top) and f (bottom) haems;

yellow, carotenoid; green, chlorophyll; ball-and-stick model represents the iron-sulfur

cluster of the high potential chain. Green lines, thylakoid membrane boundaries.

Adapted from (Malnoe et al, 2011).

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The chemiosmotic theory and ATP production

Peter Mitchell’s research that preceded the Q-cycle formulation is his work about the

chemiosmotic theory. Later awarded the Nobel prize, he proposed in 1961 that

processes leading to the production of ATP, the major energy currency of the cell,

are associated with lipid bilayers, and that the ADP phosphorylation uses an

electrochemical gradient of protons formed across the membrane.

His theory postulated that during cellular respiration in mitochondria the chemical

energy of sugars was used to transport protons through a membrane to a

compartment against the chemical gradient. It is now known that the bioenergetics of

chloroplast is similar. Because the concentration of protons would be higher in this

compartment – lumen in the case of the chloroplast - a difference in pH would be

established between lumen and stroma. Moreover, as protons are positively charged,

the driving force for the equilibration (i.e. potential energy) between these

compartments would only increase due to the electric component. Hence the

membrane would become energized and could be then defined with regards to its

positively charged side (p) facing the lumen and the negatively charged, stromal face

(n). Thanks to the fact that the thylakoid membrane is not permeable for protons, this

electro-chemical gradient can be however exploited by an enzyme synthesizing the

ATP. The ATP synthase is a molecular motor, which transforms the potential energy

of the electrochemical gradient of protons into kinetic energy of its rotor. The rotor

then interacts with the catalytic units, which drive the phosphorylation of ADP by

inorganic phosphate (fig. 7).

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Fig. 7 ATP synthesis. A, structure of ATP synthase from bovine mitochondria. The

chloroplast enzyme is similar, with spinach c subunit stoichiometry of 14. B, a

diagram of the mechanism of proton translocation. The -COO- group of an acidic

residue (usually Glu) of the c subunit is protonated by luminal H+ and the resulting

neutral residue can face the low dielectric constant lipid phase. After a full turn, it is

deprotonated and the H+ ion is released to the stroma, while the charged –COO-

facing the a subunit can continue the turn back to the starting position. C, catalytic

mechanism of the ATP synthesis by the F1 part of the enzyme driven by

conformation changes induced by the ɣ subunit. Adapted from (von Ballmoos et al,

2009) (A); and Lodish et al., Molecular Cell biology, 6th ed. (C);

As described above, PSII and cyt. b6f both contribute to the formation of pH by

releasing protons in the lumen. Furthermore, both photosystems and the b6f also add

to the electric component. Photosystems do that thanks to the charge separation

(creating positive charge in the lumen and negative on the stromal side) and the cyt.

b6f due to the fact that part of the electrons travel from the luminal (p) to the stromal

(n) side of the complex during the Q cycle.

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Fig. 8. A scheme of the energized thylakoid membrane with structures of the major

actors involved in linear electron flow (LEF) from water to NADPH. Absorption

spectra of plant PSII and PSI supercomplexes, together with characteristic absorption

peaks of the primary donors of each are shown as dashed orange and blue lines,

respectively. Adapted from (Croce & van Amerongen, 2014).

Calvin-Benson-Bassham cycle

Having generated ATP and NADPH, the chloroplast can use them to fix the inorganic

carbon and transform it into sugars. This occurs via the Calvin-Benson-Bassham

(CBB) cycle, also called the reductive pentose phosphate cycle (fig. 9). Taking place

in the stroma of the chloroplast, the CBB cycle uses ATP and NADPH in a fixed

stoichiometric ratio of 3:2 in order to allow reduction of CO2 and its transformation

into glyceraldhyde-3-phosphate, and further into sugars. The most prominent enzyme

of the cycle is the Ribulose bis-phosphate carboxylase/oxygenaze (RuBisCO), a

hexadecameric complex catalyzing an addition of inorganic carbon to a C5 molecule,

ribulose-1,5-bisphosphate. Contrary to a popular belief, this series of biochemical

reactions does not take place in the dark due to complex redox machinery regulating

it – i.e. the enzymes of the cycle are only active when reduced.

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Fig. 9. Calvin-Benson-Bassham cycle. OpenStax CNX CC BY 3.0.

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Regulation of photosynthesis

Chemistry of photosynthesis inevitably involves radicals, excited states and other

unstable, highly oxidizing or reducing compounds (Rutherford et al, 2012). It is well

known that those can be detrimental to the cell due to their reactivity. In particular,

formation of reactive oxygen species (ROS), a direct result of the presence of

radicals and molecules with highly negative redox potential can be damaging, as the

ROS readily react with organic compounds such as lipids, proteins and nucleic acids

(Schmitt et al, 2014).

A look at the Z-scheme reveals that reducing compounds are generated by light. On

the other hand, bottlenecks of photosynthesis, i.e. reactions which are kinetically

limiting the process are the b6f -mediated quinols oxidation and the activity of

RuBisCO in the CBB cycle which occur downstream of the light reactions. It is

therefore easy to understand that an excess of light – or lack of CO2 is potentially

harmful for the cell. Chloroplast, as a consequence, developed elaborated ways to

protect themselves from such imbalance aiming at an efficient regulation of

photosynthesis while maintaining a maximal possible electron flux in order to grow.

Instead of a simple scheme, because of the multitude of regulatory processes, the

thylakoid membrane can be represented in a more complete manner (fig. 10). It is

essential to think about photosynthesis “holistically” – taking into account its

structuration within the chloroplasts (structure of the thylakoids), within cells

(interactions with metabolic pathways, in particular with cellular respiration in the

mitochondria), and sometimes also tissues (in particular types of photosynthetic

organisms). Some of the most important regulatory mechanisms are described

below.

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Fig. 10. The complexity of the thylakoid membrane with selected regulatory factors.

From (Eberhard et al, 2008).

State transitions

State transitions are a reversible process regulating the cross-section of PSII and PSI

(Goldschmidt-Clermont & Bassi, 2015). It consists of a phosphorylation and then a

lateral movement of LHCII antenna from the appressed regions of thylakoid

membrane, where it formed supercomplexes with the PSII toward the non-apressed

regions. In the stroma lamellae, they increase the cross-section of PSI-LHCa

supercomplexes.

State transitions are a mechanism that balances the excitation energy between the

photosystems in light-limiting conditions when the rate of PQ reduction is higher than

its oxidation. This can occur because of an unequal excitation of the photosystems

due to differences in their absorption spectra (see fig. 8), because of an additional

non-photochemical reduction of the PQ pool or in conditions when no PSI electron

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acceptors are available. The two latter occur in particular in algae, the former in

higher plants.

This regulatory process is triggered by an overreduction of the PQ pool. It was shown

that the kinase that phosphorylates the LHCII, known as stt7 or stn7 (in algae and

higher plants, respectively) is activated when the Qo site of the cyt. b6f is occupied by

a quinol and not a quinone (Zito et al, 1999). A negative feedback from the reduced

pool, therefore, both increases the rate of its oxidation by the PSI, as well as

decreases the rate of reduction via the PSII. The STT7 kinase was shown to be

associated with the b6f, but, interestingly, neither the exact mechanism of this

sensing nor the mode of action of this kinase (i.e. how can it phosphorylate the LHCII

being itself present in the non-appressed regions) were not revealed as of now.

NPQ

Non-photochemical quenching describes two processes from an experimental

perspective, an observation that in high light photosynthetic organisms emit less

fluorescence due to an increased conversion of excited state to heat (Rochaix,

2014). The following two processes contribute to this effect: the so-called energy-

dependent quenching, called qE, and photoinhibition (qI). Although upon State

transitions a decrease of fluorescence is also observed, it cannot be described as

non-photochemical because it occurs due to a quenching by PSI.

A regulatory role of photoinhibition can be debated: it takes in fact a light-induced

PSII damage in order to observe this type of quenching. In the case where the light

intensity strongly exceeds the capacity of photosynthesis, the active center of PSII is

exposed to ROS-induced photodamage. However, upon photoinhibition the excitons

in PSII are brought quickly to the ground state, preventing further formation of singlet

oxygen (produced when the non-reactive triplet oxygen interacts with triplet states of

chlorophylls).

The qE quenching, on the other hand, is a typical regulatory mechanism. Triggered by

an acidification of the thylakoid lumen, two separate processes contribute to a

decrease of exciton lifetime in the antennae. The first is an activation of luminal

enzymes responsible for de-epoxidation of zeaxanthin, an antennae-bound

carotenoid. Its de-epoxidized form, violaxanthin, is an efficient quencher when it

accepts the excitation energy from a neighbour chlorophyll molecule. Second

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mechanism is more direct – when the concentration of hydrogen ions in the lumen

increases, they can protonate residues of the PSII antenna. This process changes

conformation of the pigment scaffold and in turn increases the probability that the

excited state returns to the ground state emitting heat.

In the case of qE, it is also a negative feedback that constitutes the protection – in

high light the electron transfer will be rapid, resulting in high rates of proton

translocation from stroma to lumen. The latter will slow down the electron extraction

from water as less excitons will end up being transferred to the special pair of PSII.

Cyclic electron flow and the „photosynthetic control“

Cyclic electron flow (CEF) is an elusive alternative electron transfer pathway. It was

discovered by Arnon in 1955 (Whatley et al, 1955), when he observed a reduction of

PQ by Flavin mononucleotide. CEF contributes to the generation of proton gradient

across the membrane through the Q-cycle without a net production of reductants.

This pathway is still largely unexplored, and many misconceptions surround it.

It is generally thought that CEF, or rather non-PSII-mediated PQ reduction operates

through two pathways (fig. 11). The minor one would consist of plastoquinone

reduction via two types of enzymes. In higher plants, a complex I-like NDH would

oxidize ferredoxin and reduce the pool, coupling this electron transfer with proton

pumping across the thylakoid membrane (Peltier et al, 2016). In microalgae, this role

would be fulfilled by a non-electrogenic peripheral thylakoid protein exposed to the

stroma, which uses NADPH as the electron donor and possess FAD as a cofactor –

the NDA2. NDA2 has also a homolog in the mitochondria, the (yeast) Ndi1 (Iwata et

al, 2012).

The second CEF route is more elusive. It is generally assumed that two proteins,

pgr5 (proton gradient regulation-5) and pgrl1 (pgr5-like 1) mediate a major – in terms

of rate - CEF pathway (Yamori & Shikanai, 2016). The mutants of those proteins,

present in both microalgae and higher plants, are impaired in the qE non-

photochemical quenching suggesting that they transfer fewer protons per light

quantum absorbed in the PSI than the WT. It is believed that they constitute a

ferrodoxin:plastoquinone oxidoreductase pathway present in the thylakoid membrane

– and that pgrl1 fulfils the role of the FQR (Hertle et al, 2013).

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Fig. 11. Proposed CEF pathways in higher plants and microalgae. Chlororespiration

(described below) is shown as it also consists of quinone reduction and oxidation.

From (Yamori & Shikanai, 2016).

There is little doubt that NDH-mediated CEF is a minor pathway. Estimated maximal

rate in vivo of this enzyme of 0,3 e-/s/PSI (Joliot & Joliot, 2002) is insufficient to

provide a significant contribution to the proton gradient (maximal steady-state LEF is

around 150 e-/s/PSI). This is despite its proposed proton-pumping role, although the

latter was never experimentally proven. Apart from very low light intensities, NDH

contribution to CEF is therefore irrelevant – but in darkness it is most probably also

the case as little reduced Ferredoxin is present in this conditions.

The pgr5/pgrl1 pathway is also surrounded by inconsistencies. It is known that the

maximal CEF rate in plants is around 130 e-/s/PSI, similarily to the maximal LEF rate

(Joliot & Johnson, 2011). It could be even hypothesized that quinol oxidation at the

Qo site is the limiting step in CEF, yet this needs to be proven. In a pgr5 mutant, the

maximal rate of CEF was not altered (Nandha et al, 2007), showing that this process

is merely regulated, and not “dependent” on pgr5. This is also the case for pgrl1,

which phenotype is identical to the pgr5 (P. Joliot, personal comm.). Those two

proteins are also highly sub-stoichiometric with regards to the photosystems and cyt.

b6f – if they indeed mediated CEF, the rate of pgr5/pgrl1-mediated PQ reduction

27

would need to be an order of magnitude higher than the maximal rate of CEF per

photosystem.

It was shown recently that CEF is regulated in higher plants at the level of FNR. The

latter can be found bound to the membrane or soluble – and interestingly co-purifies

with cyt. b6f. It has led some to propose that ferredoxin, baited by the FNR donates

electrons directly to the stromal haems of the b6f (bh or ci). Since multiple screening

campaigns, which were aimed at revealing the CEF – and not LEF – actors failed, it

is conceivable to think that CEF operates in fact as the Q-cycle initially proposed by

Peter Mitchell (fig. 12).

In microalgae, maximal rate of CEF is problematic to measure due to a much less

red-shifted PSI absorption spectrum, which largely prevents the use of far-red light as

preferentially exciting PSI. Anoxia is therefore used to induce a state of high CEF,

which is generally thought to occur because of a structuration of CEF components

(iwai; takahashi). The lack of pgr5 and pgrl1 seems to impact the rate of CEF in

anoxia, but not in oxic conditions, which would be in disagreement with the higher

plants mechanism (Alric, 2014).

Studies that address the question of actual rates of CEF in the absence of PSII

inhibitors reveal that very high rates are not achieved (Godaux et al, 2015b) even in

anoxia, and that CEF is mostly transitory.

28

Fig. 12. CEF could operate by reducing quinones at the Qi site of the b6f by

ferredoxin. From (Joliot et al, 2004b).

As for the NDA2, although theoretically it also can mediate CEF, due to its low

maximal rate of 2 e-/s/PSI (when the PQ pool is oxidized, a situation which does not

happen in the absence of PSII inhibitors) its contribution to the proton gradient is

most probably irrelevant. Moreover, the canonical CEF does not go through the

NAD(P)H pool, i.e. electrons participating in CEF return to the membrane directly

from the FNR. Because of this, NDA2 will be considered throughout as a

chlororespiratory enzyme, contributing to the redox maintenance of the chloroplast in

darkness, rather than a CEF component.

If the rapid CEF (130 e-/s/PSI in plants; 60 e-/s/PSI in algae) indeed does not pass by

the NAD(P)H pool, it is a light-induced process in vivo. This is because ferredoxin

due to its low redox potential is reduced only in the light, when its reduction is fuelled

by the PSI. Even if theoretically it is also reduced in darkness and anoxia in algae,

CEF in these conditions is not sustainable because of a lack of electron acceptor.

What is the role of CEF? Because reduction of a quinone by Fd necessarily takes

place in the stromal side of the membrane, the uptake of protons and their

29

subsequent loss upon oxidation of the quinol by the b6f is electrogenic (contributes to

an establishment of electrochemical gradient of protons across the thylakoid

membrane). Therefore, light via CEF can generate ATP without production of

NADPH (see figs 8 and 11).

This led some authors to propose that CEF compensates for a lack of ATP generated

by the linear electron flow. The latter results in 6 protons in the lumen per 2 electrons

transferred to NADP+ (a two-electron transporter). Taking into account the structure

of plant ATP synthase it was hypothesized that at least 14 protons are needed in

order to phosphorylate 3 ADP molecules (so 6 protons would yield 1,3 ATP)(Allen,

2003). As described before, the CBB cycle requires 3:2 stoichiometry between ATP

and NADPH, yet if the above reasoning is correct, only 2,6 ATP molecules would be

provided for 2 NADPH without CEF. On the other hand, it is proven that thanks to the

exchange of reductants between chloroplasts and mitochondria, lack of CEF can be

compensated by ATP synthesis from NADH in order to maintain photosynthesis

(Bailleul et al, 2015). Finally, CEF was proposed to have a role in photoprotection,

which goes two-way. As it is capable of lumen acidification it slows down the PSII

turnover via qE, but recently a novel hypothesis emerged linking it to PSI

photoprotection. The latter occurs because due to the decrease in luminal pH, the

release of protons during quinol oxidation at the Qo site of the b6f becomes

unfavourable energetically. This results in a lowered rate of b6f turnover (i.e. the rate

of the already limiting reaction further decreases), which in turn leads to an oxidation

of the PSI special pair. In case of an acceptor side limitation, CEF could hence

contribute to a slowing down of charge recombination in the PSI. Finally, P700+

(accumulated the slower works the b6f, and the higher the light intensity) is an

excellent non-photochemical excitation quencher, which would also prevent a

conversion of chlorophyll triplet states into singlet oxygen.

The pH decrease-dependent slowing down of the b6f is also known as the

photosynthetic control, and constitutes another negative feedback loop capable of

reducing eventual damage to the photosynthetic apparatus.

Oxygen as an electron sink

Oxygen can readily accept electrons from compounds that have sufficiently negative

redox potential. This is the case for primary electron acceptors in PSI (the iron-sulfur

30

clusters) and Ferredoxin, as well as NADPH. If the rate of the CBB cycle is limiting,

eventually those compounds will transfer their electrons to O2. This is the case for

Mehler reaction, in which PSI cofactors or Ferredoxin reduce molecular oxygen,

forming superoxide (O2−).

Although the Mehler reaction itself is a passive process, i.e. not a contrieved

biological regulation, the presence of superoxide dismutase and catalase – enzymes

which metabolize the superoxide to peroxide (SOD) and peroxide to water (catalase)

– suggests an adaptation to a presence of these ROS.

Flavodiiron proteins, on the other hand, are able to harmlessly reduce molecular

oxygen to water using NADPH in the chloroplast stroma. This sink for surplus

reductants is specific to more primitive organisms, like cyanobacteria (Allahverdiyeva

et al, 2013), green algae (J. Alric, in preparation) and mosses (T. Morosinotto et al.,

in press) and is crucial at the onset of illumination and in conditions of fluctuating light

– most probably before CBB cycle’s enzymes are fully activated.

Photorespiration is another process where oxygen is an electron acceptor. It is a

result of RuBisCO oxygenating instead of carboxylating its substrate. These mishits,

occurring when the concentration of CO2 is low, are costly to the cell, as each

ribulose bisphosphate molecule oxygenated requires 9 ATP molecules and 1 NADPH

molecule to regenerate in a complicated manner by chloroplast, cytoplasmic and

mitochondrial enzymes.

Finally, PTOX – plastid terminal oxidase – is an enzyme able to reduce molecular

oxygen and involved in chlororespiration, another alternative electron pathway. It will

be described in the following chapter.

31

PTOX and chlororespiration

Historical view

Early observations

It all started with the Kok effect (Kok, B. (1948) Enzymologia 13, 1-56; and (Kok,

1949)). Nearly 70 years ago, it was observed that light response curves of oxygen

evolution as a function of light intensity deviate from linearity below saturation at very

low light intensities. This was initially described as a light-dependent suppression of

respiration above the “Kok intensity” (~1000 lux).

To the knowledge of the author, the first proposal that the Kok effect was actually due

to a respiratory chain present in the chloroplasts came in 1963 from Goedheer

(Goedheer, 1963). The author showed, using luminescence, a non-photochemical

reduction of the electron carriers between the two photosynthetic units.

Subsequent studies, in particular a publication from Diner and Mauzerall (Diner &

Mauzerall, 1973) shed more light on the relation between oxygen and the chloroplast

electron transfer chain. Using repetitive saturating flashes with varying frequencies

and in condition of anoxia they demonstrated that contrary to previous hypotheses, it

is a light-independent reduction of photogenerated oxidized electron transporters in

the photosynthetic electron transport chain which is responsible for a drop in

photochemical activity when the frequency of flashes decreases. Strikingly, the

authors proposed a scheme, which is remarkably similar to the one we presented

after 40 years of research on chlororespiration (fig. 1). The major contribution of

these authors was the demonstration of a rapid rate of PQ reduction by – supposedly

- NAD(P)H. Because in oxic conditions, or when using far-red light, which

preferentially excites PSI, superimposed on flashes, the drop in photosynthetic

efficiency was markedly lower, it could be concluded that an oxidative activity, i.e. the

presence of oxidized electron acceptors, was crucial for an efficient charge

separation.

32

Fig. 1, top: from (Diner & Mauzerall, 1973) Z-Q and P-X – donor-acceptor couples in

PSII and PSI, respectively; Q - QA; A - PQ pool; R – stromal reductants, or NAD(P)H

pool. Bottom, from (Nawrocki et al, 2015).

33

Independently, Healey and Myers (Healey & Myers, 1971) showed that the Kok effect

was in fact a P700+-induced “photodepression in O2 uptake”, later re-discovered by

Peltier et al. using 16O2 and 18O2 isotopes (Peltier & Schmidt, 1991; Vermeglio et al,

1990).

In 1980, another observation confirmed that a respiratory chain must be present in

the photosynthetic membrane. The work of Godde and Trebst (Godde & Trebst,

1980) proved that in the light and presence of DCMU, if reductants were added to a

chloroplast fraction from broken cells, the hydrogen evolution in anoxia increases.

Notably, NADH addition enhanced the light-driven production of H2.

The concept of Chlororespiration

In 1982, Pierre Bennoun in IBPC coined the term Chlororespiration to differ it from

mito- and cytorespiratory activities (Bennoun, 1982), but also from photorespiration, a

process where RuBisCO oxidizes rather than carboxylates its substrate. He tried to

account for two separate observations, that of PQ being a part of a respiratory chain

(thus reduced independently of light and oxidized by oxygen), and another one, that

the thylakoid membrane was being kept energized in darkness. Logically, the free

energy from quinol oxidation by oxygen could be used to couple proton translocation

to the lumen and thus be used to generate ATP, similarily to what is happening

during mitochondrial respiration. This was the reason behind his proposal that

chlororespiration indeed stored the energy of NADPH oxidation in the proton

gradient, together with an observation that the use of cytochrome c oxidase inhibitors

resulted in a reduction of the PQ pool (suggesting a presence of a complex IV-like

oxidase in the chloroplasts). However, he has revised this view in subsequent papers

(Bennoun, 1994; Bennoun, 2001; Bennoun, 2002; Rappaport et al, 1999) where it

was proven that the loss of photochemical gradient in darkness in anoxia was not

due to suppression of chlororespiration, but mitorespiration, and that the apparent

sensitivity of the plastidial oxidase to cyt. c oxidase inhibitors was in fact due to an

export of reductants from the blocked mitorespiratory chain as well as increased

glycolysis (partially in algal chloroplasts) occurring because of a drop in ATP

concentration (Gans & Rebeille, 1990). The latter was proven with a bc1 complex

mutant which conferred resistance to myxothiazol, an otherwise specific inhibitor of

the bc1: upon addition of the drug in the mutant, the NAD(P)H and PQ pools were not

34

reduced (Bennoun, 1994). Substantiating the notions of close relations between the

two organelles in the cellular metabolism, he showed that the thylakoid membrane

lost the electrochemical gradient due to a decrease of ATP concentration (by

suppression of ATP synthase running in the proton-pumping direction).

In 1991, Peltier and Schmidt presented evidence for a regulation of the

chlororespiratory activity. During nitrogen starvation in Chlamydomonas, an increase

in oxygen signal can be seen after first two saturating flashes with regards to a

nitrogen-replete sample by an oxygen electrode (Peltier & Schmidt, 1991). This

method takes advantage of the fact that oxygen evolution occurs only after the third

flash in a dark-adapted sample (because in darkness the PSII water-splitting

manganese cluster is mostly in the S1 state (Govindjee, 1975; Joliot, 2003; Kok et al,

1970; Suga et al, 2015). The apparent increase in oxygen evolution after the first

flashes is, similarly to the one observed in continuous illumination over 1000 luxes by

Kok 40 years earlier, an actual light-induced suppression of oxygen consumption.

Peltier and Schmidt showed that the PQ pool was also more reduced than in the WT

(by the means of the FV/FM comparison), indicating that the reducing activity was

even more increased than PQ oxidation.

Molecular identification of the chlororespiratory proteins

Although already Diner and Mauzerall (1973) wondered whether it was possible that

PQH2 undergoes a direct oxidation by oxygen, it was not until the ‘90s when the first

molecular actors involved in chlororespiration were discovered.

As chloroplast genomes began to be sequenced, it was discovered that in higher

plants there were genes the sequences of which resembled those of complex I

subunits. The latter is the main entry point of reductants to the mitochondrial electron

transport chain (aside the succinate dehydrogenase) and catalyzes an oxidation of

NADH and reduction of ubiquinone, coupling it to proton translocation to the

intramembrane space of the organelle. It was therefore plausible to assume that an

NDH, NAD(P)H dehydrogenase is responsible for the R->A part of the pathway

described in the scheme above. The lack of ndh genes in the chloroplast genomes of

green algae, however, was intriguing as the non-photochemical reduction of PQ was

well documented (and discovered) in unicellular organisms.

35

In 1999, an Arabidopsis variegation mutant was described simultaneously by two

groups (Carol et al, 1999; Wu et al, 1999). Variegation is the presence of white

sectors of the leaf, which do not possess developed chloroplasts, nor pigments. The

mutant, IMMUTANS, isolated long before was shown to be impaired in carotenoid

biosynthesis; however, the mutated gene turned out to code for an oxidase which

bear sequence resemblance to the alternative oxidase (AOX) from mitochondria

(Vanlerberghe & McIntosh, 1997). As later shown, it is also a diiron oxidase. AOX

was involved in an alternative electron flow and allowed bypassing the bc1 complex

thanks to its ability of quinone oxidation. In the plant mutant, carotenoid biosynthesis,

taking place during the earliest stages of chloroplast development, could not

proceed. This is because it involves a desaturation of carotenoid precursors, a

process that necessitates oxidized quinones as electron acceptors. The lack of the

oxidase in IMMUTANS resulted in an overreduction of the pool of quinones and

directly or indirectly impeeded the development of the nascent organelle. The second

actor of chlororespiration thus had been discovered – and IMMUTANS has been

renamed as the plastid terminal oxidase, PTOX.

Final parts of the puzzle were assembled when the Peltier group resolved the

mystery surrounding the NAD(P)H:PQ oxidoreductase. It has been first proven by

Godde (1982) that a FAD/Fe-S-containing NADH:PQ oxidoreductase was present in

Chlamydomonas thylakoids. However, the discovery of the type-II NAD(P)H

dehydrogenase that is responsible for this reduction process only came in mid-2000s

(Desplats et al, 2009; Jans et al, 2008; Mus et al, 2005). Nda2, similarly to PTOX is a

non-electrogenic, monotopically bound peripheral membrane protein that uses FAD

as a cofactor.

Finally, structures of homologues of NDA2 and PTOX were described as the Ndi1

from yeast mitochondria and AOX from Trypanosoma bruceii were crystalised (Iwata

et al, 2012; Shiba et al, 2013). No structure of the photosynthetic NDH is available as

of now, and although it bears resemblance to the L-shaped complex I (Arteni et al,

2006; Baradaran et al, 2013; Kouril et al, 2014) it possesses a significant number of

subunits that the latter does not, yielding the comparisons difficult (Peltier et al,

2016).

36

Decades after the discovery of the chlororespiration process, its actors were

revealed, but the most important question remained: why would chlororespiration be

retained during evolution, if it consists of oxidation of the reducing power in a

process, which does not conserve the released energy? We tried to address this

question in a following review from the PTOX standpoint. We note here that since

then, a counterpart of our review also has been published about the NAD(P)H:PQ

oxidoreductases in the same journal (Peltier et al, 2016).

37

PTOX review

38

Annu. Rev. Plant Biol. 2015. 66:49-74

doi: 10.1146/annurev-arplant-043014-114744

Copyright © 2015 by Annual Reviews. All rights reserved

<doi>10.1146/annurev-arplant-043014-114744</doi>

Nawrocki et al.

The Plastid Terminal Oxidase

The Plastid Terminal Oxidase: Its Elusive

Function Points to Multiple Contributions to

Plastid Physiology

Wojciech J. Nawrocki,1 Nicolas J. Tourasse,2 Antoine Taly,3 Fabrice

Rappaport,1 and Francis-André Wollman1,2

1Physiologie Membranaire et Moléculaire du Chloroplaste, CNRS–Université

Pierre et Marie Curie UMR 7141; 2CNRS FRC 550; and 3Laboratoire de

Biochimie Théorique, CNRS UPR 9080, Institut de Biologie Physico-Chimique,

75005 Paris, France; email: wollman@ibpc.fr

Keywords PTOX, chlororespiration, chloroplast, electron transfer, structural

model, carotenoid biosynthesis, redox poise, retrograde signaling

■ Abstract Plastids have retained from their cyanobacterial ancestor a fragment

of the respiratory electron chain comprising an NADPH dehydrogenase and a

diiron oxidase, which sustain the so-called chlororespiration pathway. Despite its

very low turnover rates compared with photosynthetic electron flow, knocking out

the plastid terminal oxidase (PTOX) in plants or microalgae leads to severe

phenotypes that encompass developmental and growth defects together with

increased photosensitivity. On the basis of a phylogenetic and structural analysis

of the enzyme, we discuss its physiological contribution to chloroplast

metabolism, with an emphasis on its critical function in setting the redox poise of

the chloroplast stroma in darkness. The emerging picture of PTOX is that of an

enzyme at the crossroads of a variety of metabolic processes, such as, among

39

others, the regulation of cyclic electron transfer and carotenoid biosynthesis,

which have in common their dependence on the redox state of the plastoquinone

pool, which is set largely by the activity of PTOX in darkness.

INTRODUCTION

Oxidases in Energy-Transducing Membranes

Oxygen, found in its reduced form as water (2) and in its oxidized form as

gaseous molecular oxygen in the atmosphere (0), is the most abundant element

on earth. It is involved in many redox processes throughout the life realm,

prominent among which are respiration and oxygenic photosynthesis, which

respectively use oxygen as an electron acceptor and water as an electron donor.

These metabolic processes involve a membrane-borne electronic flow that is

coupled to the synthesis of ATP, thus supplying cells with the energy currency

required to sustain their metabolism, development, and division (87, 102). As a

consequence of its two unpaired electrons occupying two antibonding orbitals,

the dioxygen molecule is highly reactive and can be found in many different

chemical states and, in particular, in reactive oxygen species (ROS)—primarily

singlet oxygen (1O2), superoxide ( 2O ), and peroxide derivatives (8, 136), which

may trigger oxidative reactions in an uncontrolled manner and are potentially

harmful to their biological environment. This may account for the cellular

enzymatic arsenal involved in the active or passive detoxification of ROS or in

the scavenging of their oxygen precursor (86, 115, 117, 131).

Oxidases use dioxygen as an electron acceptor. Whereas bacteria encode a

variety of oxidases, some of which are primarily oxygen-scavenging enzymes

(50), mitochondria harbor two oxidases (for a review, see 135): the cytochrome c

oxidase (complex IV) and the alternative oxidase (AOX), first described in plant

mitochondria (12; for a recent review, see 100). Complex IV is a diheme

dicopper oxidase. It is a transmembrane, electrogenic protein complex that

couples dioxygen reduction to proton pumping across the membrane (160, 164).

A water-soluble cytochrome c found in the intermembrane space acts as its

electron donor. AOX is a simple diiron protein, bound monotopically to the n-side

of the inner mitochondrial membrane (93, 154, 166). It sustains a

40

nonelectrogenic, cyanide-insensitive (in contrast to complex IV) respiration that

uses quinols, which are soluble in the lipid bilayer, as electron donors. At first

sight, the quinol-oxidizing activity of AOX may be seen as futile from a

thermodynamic standpoint or as a partially decoupling electron flow from proton

pumping, because it lowers the ATP yield of the respiratory pathway by diverting

reducing equivalents from the sequential electrogenic activities of complexes III

and IV. However, its affinity for oxygen being lower than that of complex IV (46),

it is a poor competitor to the downstream cytochrome bc1 oxidase complexes

unless the quinone pool is overreduced (73). Thus, it may act as a safety valve

that would function only when the mitochondrial electron transfer chain is

congested. There are, however, many other possible roles for AOX that stem

from its nonelectrogenic behavior, including thermogenesis (156), oxygen

scavenging (46), stress signaling (30), reactive oxygen and nitrogen species

homeostasis (90, 154), gametophyte development (27), fruit ripening (113), or,

when working as an overflow device, a damper of metabolite fluctuations (120).

Oxidases: enzymes that reduce oxygen; they are either bioenergetic enzymes

engaged in oxidative phosphorylation or scavenger enzymes decreasing the

local oxygen concentration

Alternative oxidases: quinol oxidases contributing to an electron transfer

pathway alternative to cytochrome oxidase; mitochondria harbor a unique

oxidase of this kind named AOX

n-side: the negatively charged side of the inner membrane of a mitochondrion

after this membrane has been electrically polarized by the turnover of the

respiratory enzymes

A counterpart of this mitochondrial AOX is specifically found in plastids: the

plastid terminal oxidase (PTOX). PTOX, which is also a diiron-containing protein,

takes up electrons from plastoquinol (PQH2) to reduce oxygen via a

nonelectrogenic process. Originally called IMMUTANS, it is a nucleus-encoded,

4-helical bundle, peripheral membrane protein. Its mass varies from 40 to 50

41

kDa depending on the species. PTOX binds strongly to the stromal side of

thylakoid membranes in their nonappressed regions (83). Its expression level in

unicellular organisms and in mature tissues of vascular plants is

substoichiometric with respect to the major protein complexes of the

photosynthetic electron transfer chain, with a PTOX/photosystem II (PSII) protein

ratio in spinach of approximately 1/100 (83). Yet, as discussed below, a variety

of stresses impact its abundance. PTOX participates in a light-independent

electron transfer pathway called chlororespiration, a term coined by Bennoun

(13) more than 30 years ago by analogy with mitochondrial respiration.

Chlororespiration involves the reduction of a pool of plastoquinones (PQs), which

are soluble in the lipid phase of thylakoid membranes, by a PQ reductase and

their reoxidation by PTOX, which acts as a PQH2:oxygen oxidoreductase.

Two types of reductases have been identified. The NADH dehydrogenase-like

(NDH) enzyme, present in land plants and some unicellular algae (Streptophyta

and Nephroselmis), uses NADPH or reduced ferredoxin (163) as an electron

donor and pumps protons similarly to its mitochondrial homolog. The type II

NAD(P)H dehydrogenase-like (NDA2) enzyme, present in most unicellular algae

(89), oxidizes NADPH (57) and is a monotopically bound, nonelectrogenic

enzyme. Thus, the overall chlororespiratory process by which NADPH (or

ferredoxin) is oxidized and oxygen is reduced, is expected to be electrogenic in

plant chloroplasts but not in algae. Strangely enough, there has been no study

so far of the potential implication of these differences in electrogenicity in the

physiological control of chlororespiration between plants and NDA2-bearing

algae.

Several reviews have been devoted to PTOX (16, 38, 91, 110, 116, 130, 146),

but its role(s) and interplay with the photosynthetic function remain enigmatic.

Here, we describe some structural features of the enzyme in light of the recent

structural data obtained for its mitochondrial counterpart. We discuss its

enzymatic function and its possible physiological integration into various

metabolic pathways, taking into account, in particular, a quantitative comparison

of its activity with the redox fluxes sustained by the pathways to which it may

directly or indirectly contribute.

THE EVOLUTION, STRUCTURE, AND FUNCTION OF THE PTOX ENZYME

42

Terminal Oxidases and Their Common Ancestor

PTOX and AOX are both diiron quinol oxidases (138, 167). With the exception of

the diiron-binding motif made of four glutamic acid and two histidine residues,

protein sequences within the nonheme diiron protein family are poorly conserved

(tbbib1819, 78). Other members in this family include methane monooxygenase,

the R2 subunit of ribonucleotide reductase, ferritin, rubrerythrin, the 5-

demethoxyquinone hydrolase, and the aerobic Mg-protoporphyrin IX

monomethylester hydroxylase involved in chlorophyll biosynthesis (19, 78, 79).

As in AOX and ribonucleotide reductase, a conserved tyrosine residue is

necessary for PTOX function (41), suggesting that quinol oxidation proceeds

similarly in PTOX and AOX. The two enzymes are found in both prokaryotes and

eukaryotes. Their common ancestor—a primitive diiron-carboxylate oxidase,

perhaps an oxygen scavenger during the transition from the anoxic to the oxic

world—is of prokaryotic origin. As pointed out by McDonald & Vanlerberghe (92),

the divergence of these two oxidases occurred before the two primary

endosymbiotic events that respectively led to the emergence of the plastid and

the mitochondrion. Indeed, PTOX and AOX are phylogenetically more distant

than their respective prokaryotic and eukaryotic versions, contrary to what would

be the case if the divergence had occurred after primary endosymbiosis. PTOX

is present in cyanobacteria; green, red, and brown algae; flowering plants; and

cyanophages. Although chloroplasts evolved from the engulfment by a

eukaryotic cell of a symbiotic photosynthetic cyanobacterium, which likely had

both a cytochrome oxidase and an alternative oxidase (49), their thylakoids

display a single type of oxidase—PTOX. Assuming that the loss of cytochrome

oxidase activity in chloroplasts is an evolutionary consequence of the integration

of both the photosynthetic and respiratory metabolisms within a single eukaryotic

cell, the same logic raises the question, addressed below, of why there has been

evolutionary pressure to conserve alternative oxidases such as PTOX in the

entire photosynthetic eukaryotic lineage.

The Number of PTOX Genes

Similarly to multiple AOX copies in plants (28), some organisms have two copies

of PTOX (157). Because PTOX shares homology with AOX, they must be

discriminated to correctly assign sequences to PTOX. We designed position-

43

specific scoring matrices for PTOX and AOX that allowed the specific retrieval of

each of these oxidases from protein sequence databases (see the online

Supplemental Material; follow the Supplemental Material link from the Annual

Reviews home page at http://www.annualreviews.org). In agreement with

previous studies (22, 91, 157), we identified several organisms from various

taxonomic groups that encode two (or sometimes three) versions of the PTOX

gene (Figure 1). Although the majority of cyanobacteria and plants contain a

single PTOX gene, species carrying two PTOX copies were found in green, red,

and brown algae; diatom; and hacrobians (cryptophytes and haptophytes). The

moss Physcomitrella also contains three copies.

Figure 1 Phylogenetic tree of PTOX protein sequences. The unrooted tree was

reconstructed using the maximum likelihood method based on a matrix of

44

amino acid substitutions computed from the multiple alignment of PTOX

sequences. Nonhomologous C and N termini were excluded, and 325 sites

were used for analysis. Supplemental Figure 1 additionally shows the statistical

confidence in branching computed by approximate likelihood ratio tests.

Organisms are color coded by taxonomic group. The inset shows an

evolutionary scheme of photosynthetic organisms based on the present

classification of eukaryotes (1, 71), using the same color coding. Most

sequences were retrieved from GenBank, and the GI number is given at the end

of each organism description. For the remainder, specific symbols have been

appended to identify the source: #, data from Phytozome; @, data kindly

provided by the authors of the genome publication (108); *, data from

unpublished genomic sequence (N.J. Tourasse & Olivier Vallon, personal

communication). The scale bar is the average number of amino acid

substitutions per site. For additional details about sequence identification and

analysis, see the online Supplemental Material.

Strikingly, phylogenetic analysis based on more than 100 PTOX protein

sequences indicates that, in most cases, the multiple PTOX sequences in a

single organism do not appear to be inparalogs (as also shown in 22, 141a).

Indeed, the individual copies of PTOX in a given species can be distantly related

and scattered in the phylogenetic tree, whereas PTOX sequences from

organisms belonging to different taxonomic groups can be closely related

(Figure 1). Phylogenetic intermixing is also seen in cyanobacteria (Acaryochloris

and Oscillatoria). Thus, species encoding several PTOX proteins may have

acquired them independently from different sources, possibly by horizontal gene

transfer.

Inparalogs: paralogs in a given lineage that all evolved by gene duplications

that occurred after the speciation event that separated this lineage from

another lineage (141a)

Because PTOX genes are present in cyanophages and algal viruses, one may

hypothesize that viruses could have acted as vectors in such gene transfers.

However, the facts that (a) a majority of cyanobacteria possess a single copy of

45

PTOX and (b) the viruses sequenced to date do not bear a copy of PTOX

homologous to any of the duplicates, are not in line with this transduction

hypothesis. PTOX was likely present in the ancestor of the Archaeplastida (92),

which comprise three monophyletic groups (1, 71) where PTOX is present (22,

91). Therefore, the scattered distribution of sequences from three red and two

blue algae (Rhodophyta and Glaucophyta, respectively) in the PTOX tree

(Figure 1) is unexpected and, unless this is an artifact caused by sampling bias,

suggests an early loss of the gene and subsequent independent reacquisitions.

According to Keeling & Palmer (72), the endosymbiotic and nonendosymbiotic

horizontal transfer of plastid-targeted genes can be relatively common in algae.

Nevertheless, in a few cases, the two PTOX copies from the same species are

more closely related to each other than to PTOX proteins from other organisms,

and this reflects organism-specific PTOX duplications (22) (Figure 1). This is the

case, e.g., for maize, Eucalyptus, Physcomitrella, red algae, and green algae

belonging to the Chlorophyceae class (including Chlamydomonas), where the

duplication seems to have occurred in the ancestor of this group. In the case of

Selaginella moellendorffii and Glycine max, the copies are nearly identical, so

although they are found in different genomic contexts, they may be false

positives resulting from artifacts in genome assembly. We note that the

cyanobacterial oxidase sequences of two Synechococcus strains (and to a

lesser extent those of Prochlorococcus strains) branch remotely and may thus

represent early-diverging or fast-evolving PTOX (Figure 1).

In conclusion, PTOX has followed a complex evolutionary history, with several

independent duplication events. The phylogenomic survey of numerous plastid-

targeted proteins shows that, in fact, such genes often display a complex

evolutionary pattern involving duplications and lateral transfers (see the

supplementary material for Reference 22).

The duplication, represented in every group of photosynthetic organisms for

which the protein sequences are retrievable, begs the question of the functional

importance of these enzymes, inasmuch as they may display differential

expression patterns or responses to the environmental conditions of the two

genes (85). We discuss this further below (see The Function of PTOX in

Thylakoid Membranes).

46

AOX and PTOX: Similarities and Differences

The AtPTOX gene (originally called IMMUTANS) is distantly related to the gene

encoding the mitochondrial AOX enzyme (26, 162). The ExxEH…ExxEH–

specific motif, responsible for iron chelation, is conserved in both oxidases and

required for function (41). As discussed above, the two enzymes have a compact

four-helix scaffold that harbors the chelation site of the active transition metal

center (tbbib1819). In addition, two amphipathic helices, found in both PTOX and

AOX, dock them to the membrane (138) (Figure 2a) to a depth of approximately

8 Å.

Figure 2 A model structure of CrPTOX2. The stromal surface of the membrane

is schematically depicted as light green spheres. (a) Overview of the PTOX

dimer in the membrane. Helix numbering is from N to C terminus. (b) Major

structural differences between CrPTOX2 (blue) and TbAOX (orange; Protein

Data Bank ID 3VV9). Green sticks show the last residue before and first residue

after the loop in PTOX. The increased length of the helices next to the binding

47

pocket in AOX is shown in pink. (c) PTOX (blue) superimposed on AOX

(orange). The dimerization domain of AOX spanning the monomers is shown in

black. Because of its low sequence homology, the corresponding domain of

PTOX was not included in the model. (d) The interface of two monomers. The

colors of the helices in one monomer correspond to the colors in panel a;

helices from the other monomer are shown in transparent gray. Conserved

residues (in 93% of PTOX sequences) in the interface are shown as thick

sticks; other residues that could make contact are shown as thin sticks. The

distance between two pairs of phenylalanine residues between the monomers

does not exceed 4 Å. Note the two pairs of glutamic acid and arginine opposite

one another and the lysine opposite the tryptophan. (e) Differences in the

binding pocket between PTOX (blue) and AOX (orange). (f) Docking software

model of PQH2-binding modes in PTOX. Residues flexible during the docking

are shown as sticks. Residues orienting PQH2 with regard to the diiron active

site by hydrophobic stacking are shown as surfaces.

The structure of AOX from the parasite Trypanosoma brucei was recently

solved at atomic resolution (HYPERLINK \l "tbbib138" \o "tbbib138" 138).

Bearing 16% identity (26% similar residues) to the Chlamydomonas PTOX2

sequence in a 227-amino-acid conserved region, the crystal structure can be

used to build a structural model of PTOX, which could not be derived before

owing to the relatively weak conservation between PTOX and the other

members of the diiron-carboxylate family that have a known three-dimensional

structure (Figure 2). We used the predicted structure and docking software to

gain insight into the structural determinants of the binding site of PQH2 in

Chlamydomonas reinhardtii PTOX2, the major terminal oxidase of this green

algae (52). Because AOX crystalizes as a dimer ( HYPERLINK \l "tbbib138" \o

"tbbib138" 138), we show the enzyme in its dimeric form (Figure 2a). Owing to a

lack of sequence conservation with AOX, the N-terminal part of PTOX as well as

a loop corresponding to a PTOX-specific insertion could not be modeled.

The modeled PTOX structure differs in some instances from early structural

predictions for AOXs (7, 17, 19). As expected, the predicted scaffold binding the

diiron active center is very similar to that of TbAOX. Yet two major macroscopic

differences arise. In PTOX, helices 1 and 2 next to the entrance of the binding

48

pocket are shorter than in TbAOX (Figure 2b, pink). This narrows down the

space available for PQ. In addition, the 16-amino-acid sequence corresponding

to AtPTOX’s exon 8 is located on the side of the protein opposite the membrane

(Figure 2b, loop flanking the amino acids shown in green). Although the N-

terminal part of the PTOX protein could not be included in the model, we suggest

that it contributes to the interaction between monomers within a dimer. Indeed, it

should interact with the loop corresponding to exon 8 of AtPTOX in the other

monomer, as shown in Figure 2c, which highlights the structural proximity

between this loop in one monomer and the superposed N-terminal part of AOX

in the other. Importantly, this hypothetical interaction between two different

domains in each monomer would account for the findings that the deletion of the

loop corresponding to exon 8 of AtPTOX—possibly in proximity to the N terminus

of the second PTOX monomer—results in the destabilization of the protein (41).

Figure 2e shows the residues conserved in 93% of PTOX and AOX

sequences in GenBank. As proposed earlier (17, tbbib1819, 39), they likely play

a role in the chelation of the two iron atoms comprising the active center.

However, as shown by Shiba et al. (138), the conserved histidines are not

directly implicated in iron binding, but rather stabilize the second shell of the

active site. The TbAOX structure shows that, as first hypothesized by Young et

al. (165), two leucine residues (TbAOX L122 and L212, the former of which is

conserved in PTOX and AOX) are located next to the diiron center and orient

ubiquinol (UQH2) so that it correctly interacts with the metal center, presumably

by hydrophobic stacking. The corresponding two amino acids in CrPTOX2 are

leucine (L220; AthL135) and valine (V312; AthV226), respectively. Besides

valine, leucine or isoleucine is found in other PTOXs in place of CrPTOX V312.

In AOX, the corresponding position is almost invariably occupied by leucine or

phenylalanine. In Arabidopsis, the L135A mutation drastically decreases PTOX

activity in vitro. The increased distance between the aromatic group of the quinol

and the shorter and less bulky side chain in the mutant may possibly destabilize

quinol binding (compare with Figure 2f).

Another macroscopic difference highlighted by the comparison of the

structures is the position of the large polar residue at the entrance of the

quinone-binding pocket. In CrPTOX2, lysine 212 narrows the entry but is quite

flexible (Figure 2f). In other PTOX proteins, this position is in most cases

49

occupied by another bulky and basic residue, arginine. This is, however, at

variance with AOX, where the corresponding position (TbAOX F99) is occupied

by hydrophobic residues, mostly phenylalanine, alanine, or leucine. Moreover,

the AOX entry site is wider, as according to the model, helices 1 and 2 are

longer than they are in PTOX. The presence of a conserved, polar, and possibly

charged residue facing the lipid phase is unusual and suggests some critical

function. It may contribute to the recruitment of PQs (see below for discussion)

or anchor the enzyme at the membrane surface. Indeed, in the case of AOX,

positively charged residues—arginines—on the interface between the membrane

and proteins have been proposed to interact with the polar head groups of

phospholipids (138).

Despite this list of differences, in general, PTOX and AOX are structurally

similar, as expected from their homologous function. Yet, as we discuss below,

they are not identical from either a biochemical or enzymatic standpoint, and the

limited but significant structural differences shown by the present comparison

may constitute the structural determinants of these functional differences.

Functional Properties of PTOX as Deduced from In Vitro and Heterologous

Expression Studies

The first report of an in vitro study of PTOX came from Josse et al. (68), who

expressed the Arabidopsis protein in bacterial membranes. The reconstitution of

PTOX in liposomes, i.e., without any interference with other oxidase activity, has

been achieved only recently (167). In addition to the enzymatic studies of

heterologously expressed enzymes, the complementation of knockout

Arabidopsis thaliana lines with a site-directed PTOX mutant protein provided

insights into the residues necessary for the enzyme function (39–41).

Interestingly, these reports supported a pronounced specificity of PTOX

toward PQH2 as opposed to other quinols (68, 167). This may be a notable

difference between PTOX and AOX because AOXa1, when targeted to plastids,

can complement a delta-PTOX Arabidopsis line, suggesting that it remains

active when PQH2 substitutes for UQH2 (40; for a recent review, see 116). The

same applies when using an overexpressed AOX2 that shows dual targeting (to

chloroplast and mitochondria) despite its endogenous mitochondria-targeting

sequence. Thus, AOX can substitute for PTOX and act as a PQH2 oxidase even

50

though its affinity for UQH2 remains higher than for PQH2, as shown by the

decrease in the oxygen consumption rate in bacterial membranes upon addition

of PQH2 in excess (40). PTOX would therefore be more stringent than AOX in its

specificity toward its oxidation substrate, and it is tempting to propose that this

specificity stems at, least partly, from the location at the entrance of the quinol-

binding site of a positively charged residue in PTOX in place of a hydrophobic

one in AOX, as discussed above. A closer look at the two binding pocket

environments shows that the residues surrounding the bound PQH2 are less

hydrophobic in PTOX than in AOX (Figure 2e). This is surprising because the

methoxybenzoquinone substituents in ubiquinone (UQ) make it more polar than

PQ, suggesting an opposite trend for the substrate specificity of the two

enzymes (123). Hypothetically, though, the hydrophobic interaction between the

lipid phase and the isoprenoid chain may not be sufficient to pull the polar head

of the UQ out of the binding pocket, thus preventing PTOX from using UQ as a

substrate (68, 167).

The oligomerization state of the enzyme is another structural issue that

deserves attention. Yu et al. (167) recently reported that purification of a

heterologously expressed PTOX–maltose-binding protein fusion yields mainly a

tetramer and to a lesser extent a dimer and a monomer. This stands in stark

contrast to data from Fu et al. (40), who reported that PTOX is uniquely found as

a monomer in nondenaturing biochemical analysis. Yet PTOX dimers have been

found in transgenic tobacco plants overexpressing Chlamydomonas PTOX1 (2).

Notably, the cysteine residues involved in dimerization and, consequently, in the

regulation of the activity of AOX (41, 153) are absent in PTOX. Yet the

dimerization may, as mentioned above, involve the N-terminus part of one

monomer and the loop corresponding to exon 8 of the other one (Figure 2b,c).

Even though it is not required from a mechanistic standpoint (see below),

dimerization may take part in the regulation and/or the (de)stabilization of the

protein. Finally, one could envisage the formation of heterodimers in species that

express several isoforms of PTOX, but this issue has not been addressed.

According to Moore et al. (93), quinol oxidation into semiquinone by these

diiron oxidases is initiated by a superoxo-diiron center upon oxidation of the

diferrous state by O2. After the release of a water molecule, the peroxo-bridged

diiron oxidizes the tyrosine residue (Tyr220 of CrePTOX2) in proximity to the

51

dimetal center; this radical then oxidizes the semiquinone to eventually produce

a quinone and the resting state, oxo-diiron (3+). The latter subsequently oxidizes

another quinol into a quinone, and this closes the catalytic cycle by regenerating

the diferrous center. Notably, the proposed mechanism involves no high-valence

iron intermediates, but does involve radical transients such as tyrosyl [as in

ribonucleotide reductase (76)] and semiquinone. Importantly, according to the

present views, these two radicals are formed during the reaction sequence

initiated by the binding of oxygen to the diferrous center and terminated by the

formation of the oxo-diiron state. They are thus transient and thought to be short

lived, thereby minimizing the production of ROS— quite a chemical feat, given

the need to couple two reactions that sequentially donate two electrons to a

reaction that requires four electrons (93).

There are, however, conditions where PTOX seems unable to take up this

mechanistic challenge. When overexpressed in tobacco lines or when having to

cohabit in the thylakoid membrane with a heterologously expressed bacterial

phytoene desaturase, PTOX is a strong ROS producer (42, 51). Interestingly, in

vitro studies recently provided further insights into the balance between the

antioxidant (oxygen-reducing) and prooxidant (ROS-producing) activity of PTOX.

At basic pH, increasing the concentration of the electron donor increased the

prooxidant activity, whereas the opposite applied at acidic pH (167).

THE FUNCTION OF PTOX IN THYLAKOID MEMBRANES

Chlororespiration

Because PTOX, by oxidizing PQH2, potentially diverts reducing power from the

photosynthetic electron transfer chain, it may interfere in vivo with photosynthetic

electron transport and CO2 fixation in the Calvin-Benson-Bassham cycle. Yet, as

discussed above, PTOX is present in the thylakoid membranes of the vast

majority of photosynthetic organisms, and the mere fact that it has been retained

throughout evolution suggests that, although possibly detrimental in some

instances, it fulfills beneficial functions. Contrary to the original proposal (14),

chlororespiration as such is not electrogenic in green algae in darkness (15,

118), as neither NDA2 nor PTOX is a proton-pumping enzyme (Figure 3d).

However, chlororespiratory pathway in organisms possessing NDH can probably

52

generate +Hμ in the dark. In the light, irrespective of their intrinsic

electrogenicity, chlororespiratory enzymes can contribute to +Hμ when coupled

to the activity of either photosystem: NDA2 can do so by sustaining the turnover

of the cytochrome b6f complex and the reduction of 700P when PSII is poorly

active or inactivated (an extreme experimental case being under PSII-inhibited

conditions) (61), and PTOX can do so when involved in an electrogenic water–

water cycle together with PSII (24). In a marine alga (Ostreococcus) and a

cyanobacterium (Synechococcus) that have low PSI/PSII ratios, likely stemming

from their adaptation to iron-limited conditions in the ocean, electrons from PSII

are redirected toward oxygen via PTOX. Notably, however, such a cycle

provides less ATP per oxidized H2O molecule than does linear electron flow

(LEF) from H2O to NADP+. In Ostreococcus, this water–water cycle accounts for

up to 30% of the total electron flow in these conditions. This is commensurate

with the estimates obtained in Thellungiella, a halotolerant relative of

Arabidopsis, which when salt stressed can also use PTOX as a safety valve to

redirect PSII-generated electrons to oxygen (142) by upregulating PTOX

fourfold. We note, incidentally, that a number of species with duplicated PTOX

are halotolerant or can cope with high salt stress (Figure 1). Moreover,

lodgepole pine has attracted attention for its enhanced oxygen-dependent

electron flow, particularly in winter (133), but contributions from the Mehler

reaction (O2 reduction at the level of PSI) were not ruled out.

+Hμ : the transmembrane electrochemical potential difference for H+

P700: a dimer of chlorophyll in photosystem I, which is oxidized ( 700P ) following

charge separation and reduced by plastocyanin

Water–water cycle: a redox cycle with water acting as the electron donor and

oxygen as the electron acceptor, thereby producing water

53

Figure 3 Plastoquinone (PQ) pool central to photosynthesis. In photosynthetic

organisms, two major modes of electron transfer can be distinguished: linear

electron flow (LEF) and cyclic electron flow (CEF). LEF involves the

photoinduced extraction of electrons from water and reduction of PQ by

photosystem II (PSII), plastoquinol (PQH2) oxidation and plastocyanin (PC)

reduction by cytochrome b6f, PC shuttling in the lumen and reduction of

photooxidized photosystem I (PSI), and light-induced electron transfer from

PSI to NADP+ via ferredoxin (Fd) and ferredoxin-NADP+ oxidoreductase (FNR).

CEF, which is light dependent, consists of recycling electrons from the stromal

side back to the PQ pool (29, 144). The PQ pool is depicted as an amphora (81).

Red arrows indicate PQ-reducing pathways, and blue arrows indicate PQH2-

oxidizing pathways. Cyan indicates enzymes, rates, or reactions specific to

eukaryotic algae; pink indicates enzymes and rates specific to land plants; and

green indicates enzymes common to plants and algae. “PMF” highlights the

fact that the turnover of the enzyme(s) contributes to the buildup of the proton-

motive force. (a) Major PQ-related redox pathways in the light. (b) PQ-related

redox pathways in the dark. The redox steady state of the PQ pool is shown for

54

dark-adapted organisms. (c) The carotenoid biosynthesis pathway in plants. (d)

A consensus view on the major reducing-equivalents-exchange pathways in

the chloroplast. The maximal rates of electron transfer (in units of electrons per

second per photosystem) are shown as measured in vivo under oxic,

nonstress conditions. The positions of the enzymes in thylakoid membranes

and locations of NDH-pumped protons are shown with respect to the stromal

(S) and lumenal (L) side. Black arrows indicate electron transport not directly

involving quinones, and gray dashed arrows indicate transport via the

malate/oxaloacetate shuttle and other pathways of reducing power exchange

with mitochondria and cytosol. For clarity, the quinone-reducing activity of

cytochrome b6f fueling the Q cycle is not shown. For the PDS and ZDS

proteins, arbitrary shapes and positions in the membrane were chosen based

on References 132, 145, and 151. Note that in algae, the more negative overall

redox balance is due to mixotrophy and the idiosyncratic localization of the

upper part of glycolysis in the chloroplast (extensively reviewed in References

61 and 62). Additional abbreviations: Fdred, reduced ferredoxin; PCred, reduced

plastocyanin.

Because electron flow, ATP synthesis, and the buildup of a proton-motive

force are so intricately coupled, PTOX must serve multiple functions as soon as

it is engaged in a water–water cycle with PSII, and considering it merely as a

safety valve seems excessively restrictive. In line with this, the role of cyclic

electron flow (CEF) has been recently proposed to go beyond that of a supplier

of +Hμ and to extend to photoprotection owing to the photosynthetic control

exerted at the level of the cytochrome b6f complex upon acidification of the

lumen (for reviews, see 64, 139). When involved in a water–water cycle, PTOX

could be seen as a part of any photoprotective processes triggered by the built-

up +Hμ across the membrane. These include the activation of the violaxanthin

de-epoxidase and the consecutive onset of nonphotochemical quenching,

photosynthetic control, etc. (43). In plants, where chlororespiration is

electrogenic, it may also play a significant role in sustaining the polarization state

of the thylakoid membrane in the dark (Figure 3b,d). The upkeep of such a +Hμ

across the thylakoid membrane is well documented (for Chlamydomonas, see

35; for vascular plants, see 67; for a review, see 77) and may serve all

55

processes requiring a proton-motive force, such as, to mention a few, protein

import into the lumen (149), activation of the CFoF1 ATP synthase (44), and pH-

dependent nonphotochemical quenching (95, 129). Thus, in NDH-containing

organisms, chlororespiration may contribute to polarizing the thylakoid

membranes and thereby indirectly take part to the various +Hμ -dependent

processes. In the case of pH-dependent nonphotochemical quenching or

activation of the CFoF1 ATP synthase, the latter of which requires the +Hμ to

reach values exceeding specific thresholds, chlororespiration would act as a

primer. It would set a polarization state at values remaining below, but close to,

these thresholds, thereby sensitizing the activation processes. Importantly, the

upkeep of this permanent +Hμ in the dark would develop at the expense of

reducing power, whereas in organisms with nonelectrogenic NDA in place of

NDH, it would rely on ATP consumption (118).

The Electron Sink Hypothesis

Because the enzymology of PTOX in vitro remained poorly characterized until

recently (167), most of our knowledge of the activity of PTOX comes from in vivo

studies. In vivo assays based on fluorescence methods (52, 80, 152, 158)

showed that the maximum rate of light-independent PQH2 oxidation varies

among species, reaching up to 5 electrons transferred to oxygen per second per

PSII center (e s1 PSII1) in Chlamydomonas (two PTOX enzymes exist in this

algae, PTOX2 being the major oxidase, with a rate of 4.5 e s1 PSII1) (52)

(Figure 3d). These values were obtained in mutants lacking cytochrome b6f

complexes to ensure the full reduction of the PQ pool upon illumination and

thereby allow the study of its light-independent reoxidation. In the case of

vascular plants, the assessment of PTOX activity does not require the absence

of cytochrome b6f because the inactivation of CO2 fixation in the dark creates the

bottleneck required for the full reduction of the PQ pool. The PTOX-mediated

oxidation rate in plants is much lower than in Chlamydomonas, reaching only 0.3

e s1 PSII1 in tomato (152) and 0.4 e s1 PSII1 in Arabidopsis (65). However,

the alpine plant Ranunculus glacialis exhibits a threefold-higher PTOX

abundance when grown in high light than when grown in low light (80). This is

mirrored at the functional level by a Vmax of light-independent PQH2 oxidation in

56

high-light-grown plants of 1.3 e s1 PSII1, three fold larger than in shade-grown

plants. In line with the above discussion, this light-dependent regulation of PTOX

activity led Streb et al. (143) to propose that, by acting as an additional electron

exit pathway, PTOX could provide photoprotection to the reaction centers.

Several authors have proposed the hypothesis that PTOX acts as a safety

valve in plants (26, 103, 107). However, the PTOX-driven PQ oxidation rates do

not exceed a few electrons per second per PSII, and this seems far from

sufficient to provide an efficient safety valve against high excitation pressure, a

situation encountered when the excitonic influx at the level of the photosystems

overcomes the chemical outflux of the CO2 assimilation pathway: Under

saturating light conditions, the maximal rate of photosynthesis—and thus the

reduction of PQ or the oxidation of PQH2, the rates of each step in the overall

process being equal by definition—is approximately two orders of magnitude

larger (~150 e s1 PSII1). This is illustrated by an early study showing that the

electron flow through PTOX decreases at the onset of illumination (111). This

decrease stems from the drain of reducing equivalents from quinol by the

oxidized cytochrome f, highlighting the competition between the speedy light-

driven electron transfer and the sluggish PTOX pathway. Because it is a poor

competitor of the photosynthetic mainstream, PTOX is unlikely to be a major

player in photoprotection, at least under constant and saturating light conditions.

Another feature of PTOX that argues against the safety valve function is its

contribution to ROS production (34, 42, 51, 167), which may damage the

photosynthetic apparatus. However, according to Li et al. (85), PTOX may

consume up to 10% of photochemically produced oxygen via the astaxanthin

biosynthesis pathway, which would lower the oxygen partial pressure in the cell

and consequently decrease ROS production (85). One could also argue that

PTOX contributes to PSII photoprotection by shifting the site of ROS production

from the appressed membranes, where the majority of PSII resides, to the

nonappressed membranes, where PTOX would be located (58, 83). This

hypothesis must be qualified, though, by the limited diffusion of PQH2 along

thylakoid membranes caused by molecular crowding, demonstrated by Lavergne

& Joliot (82; for a recent review, see 75). Thus, under sustained photosynthetic

electron flow at saturating light intensity, those PQs that are reduced by PSII in

57

the appressed membranes would only rarely be reoxidized by a remote PTOX

enzyme located in the nonappressed membranes.

On the one hand, the abundance of PTOX seems to be positively correlated to

abiotic stresses such as cold, high light, and high salt (55); on the other hand,

however, the artifactual overexpression of PTOX did not provide additional

protection in cold-stressed, cold-adapted, or control Arabidopsis plants (128).

These seemingly conflicting observations may be reconciled if, in natural

conditions, the increased abundance of PTOX is coupled to the upregulation of

enzymes such as superoxide dismutase. PTOX, as a link in this detoxifying

pathway, would contribute to lowering the yield of ROS production by providing a

safer way out for the reducing power produced in excess by PSII. However, such

a concerted and complementary pas de deux will not take place when a strong

constitutive promoter drives PTOX overexpression, unless ROS or PTOX directly

functions as a trans-acting regulatory element on the genes coding for

detoxifying enzymes, which does not seem to be the case (51, 58).

At this stage, an excess of PTOX under low-light conditions may be as

detrimental as (or even more detrimental than) a low amount of PTOX under

high light. PTOX activity may even generate more ROS per enzyme turnover in

low light if, as substantiated by Feilke et al. (34), the stepwise delivery of the two

quinols lengthens the lifetime of reactive intermediates so that ROS production

increases with the time delay between flashes. Thus, ROS production by PTOX,

which is not an asset for a safety valve, should probably be considered from

another perspective: It could be a starting point for redox signaling within the cell

before the PQ pool becomes overreduced, as shown for singlet oxygen

production in other physiological circumstances (8, 74).

The Carotenoid Biosynthesis Hypothesis

In contrast to the elusive role of PTOX in fully grown plants, there is little doubt

that it plays a role in chloroplast development. immutans mutants of Arabidopsis,

eventually characterized as lacking PTOX (26, 162), were obtained years ago by

random mutagenesis (122). The recessive mutation resulted in variegated

leaves, the macroscopic and visual signature of undeveloped chloroplasts, in

which neither carotenoids nor chlorophyll was synthesized. Interestingly, only

some of the chloroplasts within a given leaf were colorless, with the rest

58

displaying wild-type characteristics, and there was no intermediate organelle

state between these two extremes. This phenotype was correlated to the light

intensity, as shown later by Wetzel et al. (159) and Ross et al. (127), because

plants grown in low light developed leaves similar to wild-type plants.

The stage at which PTOX activity is critical for chloroplast biogenesis has

been studied by examining the expression pattern of the ptox gene. Notably,

only a few percent of the wild-type content of PTOX is sufficient to warrant the

proper development of leaves tissues (39). Although in Arabidopsis its

expression hardly changes in response to most environmental stresses (128), it

is regulated during plant development, and its pattern strongly supports its

involvement in early plastid biogenesis. Moreover, as shown by Wetzel et al.

(159), in PTOX-less plants, high light is harmful to developing chloroplasts only

during the 24 h following seed coat breakage, i.e., during the differentiation of

proplastids into chloroplasts (see 38 and references therein).

Cell imaging recently showed that young leaf primordia of PTOX-less

Arabidopsis in high light present a severely reduced number of chloroplasts,

further supporting the involvement of PTOX in the development of the leaf tissue

(38). Thus, PTOX activity is critical at an early stage of chloroplast development,

but its molecular action in this respect remains a matter of debate. The

mainstream view is that, in the absence of light-driven oxidation of PQH2, the

oxidase activity of PTOX provides the electron-accepting substrate of phytoene

desaturase (Figure 3d), which, in the carotenoid biosynthesis pathway (Figure

3c), catalyzes the conversion of phytoene to zeta-carotene using PQ as the

electron acceptor (32, 45, 132, 140). In the absence of PTOX, the PQ pool is

fully reduced owing to the activity of PSII in the light or that of the

dehydrogenase in the dark, and this blocks the carotenoid biosynthetic pathway

at its very early steps. (159). Consistent with this assumption, the rate of

carotenoid synthesis is markedly lower in an etiolated PTOX-less Arabidopsis

(25). This negatively impacts photoprotection, thereby inducing photooxidative

stress and preventing the proper maturation of developing chloroplasts. One

should keep in mind, however, that any process that depends on the redox state

of the PQ pool and is critical for chloroplast development, such as chloroplast-to-

nucleus retrograde signaling (37, 104), would be similarly affected in the lack of

PTOX (Figure 3d).

59

Even though the carotenoid biosynthesis hypothesis straightforwardly relates

the lack of PTOX to a defect in chloroplast biogenesis, it fails to account for the

fact that some plastids apparently cope with the lack of PQ oxidase, as

evidenced by the variegated phenotype. This latter observation is likely the origin

of the hypothesis that a threshold exists above which both carotenoid

biosynthesis and plastid development take place (159; for a recent review, see

38). This idea is consistent with the observations that (a) healthy leaves develop

in PTOX-lacking plants when grown in low light (6, 38, 122, 127, 128, 159); (b)

variegation becomes more pronounced at lower temperatures, when the light-

induced PQH2 oxidation is hampered by the lower turnover rate of the Calvin-

Benson-Bassham cycle (127); and (c) de-etiolation is similarly impaired in high

light in a PTOX-lacking strain, in which cotyledons develop variegation and the

kinetics of the rise in PSII photochemical efficiency develop more slowly and to a

lesser extent than in the wild type (127). Each of these findings is readily

accounted for if, at the early stage of chloroplast development, the stammering

photosynthetic electron transfer chain is sufficient to ensure a level of oxidized

PQ compatible with carotenoid biosynthesis, at least under low-light conditions.

When in excess, illumination would increase the reducing pressure, and PTOX

would then become a major player determining the redox state of the PQ pool.

As development would go on, the light-driven electron flux would take over, and

PTOX would be relegated to other physiological roles.

Notably, by providing an electron sink downstream of the PQ pool, PTOX

could contribute to the photoprotection of the nascent organelle in other ways

than merely supporting the carotenoid biosynthesis pathway. For instance, an

analysis of greening in etiolated pea (Pisum sativum) shoots showed that both

PTOX and the NdhI subunit of the NDH complex were present in dark-grown

plants (83). During de-etiolation, PSII subunits were more abundant than PSI

subunits after 6 h of greening (70). Therefore, as in the Ostreococcus and

Synechococcus cases described above (10, 24), basal PTOX levels in

etioplasts, increasing rapidly upon transition to light (83), may partly compensate

this unbalanced stoichiometric ratio and thereby contribute to photoprotection.

Importantly, such a hypothesis requires that the activity of the NDH complex

does not counteract the oxidation of PQ by PTOX, which seems likely given that

its substrates (reduced ferredoxin and/or NADPH) are provided mainly by PSI

60

activity. Conversely, in etiolated barley, where PSI activity would precede PSII

activation upon transition to the light (105), PTOX could act as a regulator of

CEF before a shift to the linear regime in the developed organ (see Figure 3a,d).

Testing the above hypothesis will require decoupling the indirect involvement

of PTOX in carotenoid biosynthesis from its role as an electron sink. Galzerano

et al. (42) took a first step in this direction by expressing in wild-type Arabidopsis

a bacterial phytoene desaturase (CRTI; see Figure 3d) that does not use PQ for

either phytoene oxidation or zeta-carotene oxidation (132). Unexpectedly, this

led to increased PTOX activity, an increased sensitivity of the ROS production to

propyl-gallate (an inhibitor of PTOX), and reduced growth rates. The next step

will be to overexpress CRTI in a PTOX-defective background, but this remains to

be undertaken.

The biosynthesis of carotenoids has also been investigated from a PTOX

perspective in algae. Obviously, issues related to chloroplast development do

not hold in algae, in which the single chloroplast divides in a mitotic manner with

a division furrow halving the chloroplast (48). After each cell division, a

significant light-dependent PQH2-oxidizing activity segregates in the mitotic

progeny, making PTOX less essential for providing the required electron

acceptors for the PDS protein. In fact, carotenoids accumulate in

Chlamydomonas mostly during the first part of the photoperiod. The synthesis

rates during the dark period are approximately two orders of magnitude lower

than in the light, and the absolute quantities of carotenoids hardly change during

the dark part of the cycle (56). Accordingly, strains lacking PTOX would produce

only ~2.5% less carotenoids when grown in a 12-h–12-h dark-light cycle, and

their pigment content would barely differ from that of the wild type, as reported

for a Chlamydomonas mutant lacking PTOX2 (52). Perhaps less expectedly, the

mutant displayed no change in carotenoid accumulation when grown in

heterotrophic conditions (acetate and darkness), i.e., in the absence of light-

driven oxidation of the PQ pool (52). The presence of PTOX1, another PTOX

paralog in this organism (see Terminal Oxidases and Their Common Ancestor,

above), operating at a rate approximately tenfold slower than that of PTOX2,

may account for this observation. In this respect, it is worth pointing out that

PTOX1 and PTOX2 differ from an enzymological standpoint, the latter being

more sensitive to propyl-gallate than the former (52), raising the possibility that

61

PTOX1 is specifically devoted to the carotenoid biosynthesis pathway. Along

these lines, in the Chlorophycea Haematococcus pluvialis, the PTOX1-specific

transcription pattern follows the carotenoid biosynthesis time course, supporting

its assignment to the synthesis of astaxanthin (157).

PTOX Activity in Darkness

The redox steady state of the PQ pool, which is poised in the dark by the

chlororespiratory pathway, differs between algae and vascular plants (Figure

3b,d). In the latter, after a long period of darkness, the pool is almost completely

oxidized [but see reports indicating that 20% of the PQ pool remains reduced in

darkness in tomato (152)]. In Chlamydomonas reinhardtii, 30% of the PQ pool

remains in its reduced state in darkness when the algae are kept under vigorous

stirring (52). This contrast may simply reflect differences in the light-independent

electron influx, which themselves may stem from different metabolite exchange

rates between the different cellular compartments. In algae, starch degradation

and the upper part of glycolysis (glucose to glyceraldehyde-3-phosphate) takes

place exclusively in the chloroplast, whereas in vascular plants it is cytosolic

(114). According to Johnson & Alric (61), the reductive flux in darkness in the

chloroplast is larger in algae (Figure 3), which may reflect the fact that starch is

the major source of reduced carbon in algal cells, in contrast to vascular plants,

which can metabolize sucrose in the cytosol. Alternatively, the differences in

chlororespiratory fluxes between green algae and vascular plants may be a

consequence of chlororespiration being electrogenic in the latter and not in the

former. The fact that NDH has to work against the permanent +Hμ it contributes

to generating may be a limitation to its turnover rate under steady-state

conditions.

Upper part of glycolysis: conversion of glucose, glucose-1-phosphate, or

glucose-6-phosphate to glyceraldehyde-3-phosphate, which in algae is

specifically located in the chloroplast

In any event, by enabling electron transfer in the dark from NADPH to oxygen,

chlororespiration links the redox steady state of the PQ pool to that of the

stroma, which is linked to the overall cellular redox poise by metabolite

62

exchanges. As an example of such a metabolic entanglement, PTOX has been

proposed in algae to act as a carbon sink that adjusts the intracellular N/C ratio

in conditions of unbalanced availability (112, 158). Indeed, the chlororespiration

rate is enhanced in Chlamydomonas during nitrogen starvation. In these

conditions, there is a profound remodeling of the photosynthetic apparatus

together with a fourfold increase in PTOX and NDA2 accumulation levels (21,

112, 158). Accordingly, chlororespiration becomes more active during nitrogen

deprivation [an increase from 1.4 to 3.4 e s1 PSII1; calculations are from data

in Reference 158].

Other Functional Hypotheses Regarding PTOX

Other physiological responses that may involve PTOX are the process known as

state transition and the switch between CEF and LEF. In plants and algae, state

transitions are changes in the relative absorption cross section of each

photosystem caused by the lateral migration of light-harvesting complex II

(LHCII) between stacked and unstacked regions of the thylakoid membranes

(reviewed in 125, 161). They are controlled by an LHCII kinase, the reversible

activation of which depends on the occupancy of the Qo site of cytochrome b6f

complexes by a PQH2 (155, 168). The lateral migration of LHCII toward PSI is

triggered by a reduction of the PQ pool, thereby decreasing the reducing

pressure exerted by PSII and increasing the oxidizing pressure resulting from

PSI turnover (31). Chlororespiration’s control over the redox state of the PQ pool

in darkness makes PTOX a key enzyme in the bioenergetics of the

photosynthetic cell because its activity is expected to set the appropriate light-

harvesting balance. Moreover, PTOX, by acting as a sensor of the redox and

ATP status in the chloroplast stroma, helps to trigger the ultrastructural changes

undergone by the thylakoid in response to changes in the energetic status of the

entire cell (20, 161).

State transitions: changes in the distribution of light energy between the two

photosystems in response to changes in the redox state of the PQ pool

When deprived of ATP, algal cells undergo an activation of glycolysis, known

as the Pasteur effect, that releases NADPH in the chloroplast stroma (121), thus

63

eliciting a reduction of the PQ pool, activation of the LHCII kinase, and eventually

an increase in PSI antenna size relative to that of PSII. These conditions are well

suited, at least in nonsaturating light conditions, to favor CEF around PSI, a

mode enabling the production of ATP that is not committed to carbon fixation,

and thus contributes to restoring suitable intracellular ATP levels in these ATP-

depleted cells (20). Subsequent studies on the relationship between state II

conditions and CEF in Chlamydomonas added support to the view that a state I–

state II transition may correspond to a switch from LEF to CEF (33, 36, 88).

However, further studies showed that the metabolic conditions required for

transition to state II (i.e., reducing conditions in the chloroplast stroma), rather

than the lateral redistribution of LHCII per se, were responsible for the

enhancement of CEF in Chlamydomonas (147), making the possible role of

PTOX in CEF even more direct. Indeed, by oxidizing NADPH in the dark and

thereby allowing the redox equilibration between the stroma and the thylakoid

membrane, chlororespiration in general and PTOX in particular regulate the

partition between LEF and CEF at the onset of illumination (152). PTOX could

also regulate CEF directly by preventing overreduction of the pool, but the

degree of its direct involvement in CEF remains to be specified.

As an illustration of the complex interplay between chlororespiration and CEF,

the inactivation of a protein thought to contribute to CEF, PGR5 (9, tbbib9596,

98 PTOX Arabidopsis (106). The

deletion of NDH PTOX context also rescues this phenotype, albeit to a

lesser extent, likely because of its less prominent contribution to CEF (106).

Importantly, PTOX, similarly to the major CEF actors (PSI and cytochrome b6f),

is localized in nonappressed membranes (3, 60, 83, 101); this favors its

involvement in CEF if, as proposed by various authors (33, 66), the segregation

of PSII and PSI between the appressed and nonappressed membranes is an

ultrastructural requirement for efficient CEF.

PTOX AS AN ENVIRONMENT-RESPONSIVE ENZYME

If, as discussed above, PTOX is indeed involved either directly or indirectly in a

variety of physiological functions (carotenoid biosynthesis, photoprotection of the

nascent electron transfer chain, redox poise, etc.), it would be unsurprising that

64

its abundance varies with environmental conditions. Both the protein and

transcript level of PTOX respond to environmental conditions (reviewed in 146),

but in Arabidopsis, its transcript level is rather insensitive to a wide range of

stresses (128). This initially led to the conclusion that its activity has minimal

influence in mature plants. However, PTOX has since been shown to be

upregulated in response to high-light treatment—sometimes coupled to other

stresses—in a variety of other organisms (47, 54, 80, tbbib9697, 109, 152, 157).

In addition, it is retained in the genomes of high-light- but not low-light-ecotype

cyanobacteria (91). Changes in the oxidase abundance in response to chilling

have also been documented (23, 55, 133). As mentioned above, PTOX is

differentially expressed during nitrogen or phosphate starvation in

Chlamydomonas (94, 112, 158), during drought in Coffea (141), and during salt

stress in halophilic Thellungiella (142). It is notable that in each of these

conditions the PQ pool potentially undergoes overreduction, either because of

the excessive influx of light-generated reducing equivalents or because of the

insufficient outflux hampered by the stress-induced downregulation of the Calvin-

Benson-Bassham cycle (53).

Importantly, in many of these cases, PTOX is not the only enzyme whose

abundance changes. Its chlororespiration partner (either NDA2 or NDH) is often

similarly regulated, which suggests that what matters is the acclimation of the

chlororespiratory flux as a whole rather than the oxidase activity per se. In stress

conditions, chlororespiratory enzymes would be upregulated to cope with the

increasing reducing pressure in the chloroplasts in the dark, and this would

provide a redox buffer, thereby smoothing the sudden changes induced by the

onset of the light-driven electron flow following illumination. Moreover, this could

indirectly play a photoprotective role by setting the redox poise so as to promote

CEF and the ensuing photoprotective mechanisms (53, 59, 64, 124, 130).

PTOX IN NONGREEN PLASTIDS

PTOX’s role in poising the redox state of the stroma, combined with the fact that

the chloroplast is the site of several biosynthetic pathways involving electron

redox reactions, makes it an obligate actor in several plant cell processes

beyond those discussed above. Recent studies have implicated it in

65

strigolactone metabolism in rice (148). Strigolactone, a recently discovered plant

hormone, derives from carotenoids and plays an important role in root

biogenesis and shoot branching (119, 137). Both rice and Arabidopsis PTOX

plants are more branched, similarly to other mutants defective in strigolactone

response (148). Notably, the interplay between chlororespiration and

strigolactone biosynthesis may go beyond the fact that they are both linked to

carotenoid biosynthesis, because Arabidopsis mutants defective in NDH are less

strongly affected in carotenoid biosynthesis but still show defects in strigolactone

biosynthesis. Indeed, Roose et al. (126) showed that a lack of one of the NDH-

related proteins results in different branching of the roots together with a slightly

altered PQ redox state. Interestingly, PTOX mRNA is also present in nongreen

tissues such as roots in abundances commensurate to those in photosynthetic

tissues (5). The PTOX promoter operates in several tissues and at various

developmental stages, with no obvious correlation to the tissue’s carotenoid

content (5). Finally, apart from PTOX, at least some of the enzymes of the

carotenoid biosynthesis pathway are present in the nongreen tissues of plants

(84), supporting their role in strigolactone biosynthesis and not simply in

carotenoid production for photoprotective purposes.

Not only chloroplasts (variegation) and amyloplasts are affected in mutants of

either chlororespiratory enzymes in higher plants. Chromoplasts, another

functional type of plastids, accumulate high amounts of carotenoids. Consistent

with their implication in the carotenoid biosynthesis pathway, the genes coding

for PTOX, PDS, and ZDS are upregulated in tomato and pepper fruits, which

display increased carotenoid levels (69). As expected, the carotenoid content of

chromoplasts is strongly affected in plants lacking PTOX activity (11). The

accumulation of pigments is also impaired when NDH is not functional (99).

Because carotenoid biosynthesis is an energy-consuming process, this may

reflect the need for a permanent +Hμ to sustain this biosynthetic pathway and

the involvement of chlororespiration in its generation.

CONCLUDING REMARKS AND RESEARCH FRONTIERS

The unquestionable piece of data regarding PTOX is its involvement in the

oxidation of the PQ pool. Thus, together with NDH/NDA2, it sets the redox poise

66

in the dark not only of the PQ pool but also of the entire stroma. This is expected

to impact, to different degrees, all the processes that either involve redox

reactions or are redox controlled. There are many such processes, of which CO2

fixation and CEF are the most obvious because they rely on the function of

protein complexes found in the same membrane as PTOX. Because these two

processes are driven by light and have turnover rates that are much larger than

the chlororespiration flux, their interplay with chlororespiration is more likely to be

transient and to become particularly important during transitions between

different light intensity regimes, the most extreme case being dark to light. The

other, light-independent processes are biosynthetic pathways, prominent among

which is carotenoid biosynthesis, but the recent results on strigolactone suggest

that the implication of chlororespiration may extend beyond that. Between these

two extremes stands retrograde signaling, which is commonly thought to be

triggered by the redox status of the chloroplast. The picture that emerges for

PTOX is that of a housekeeping function. It sets the appropriate redox poise and

electron transfer flow in the dark to directly or indirectly allow several other vital

functions to take place smoothly.

SUMMARY POINTS

1. The plastid terminal oxidase (PTOX) has a complex evolutionary history, with

several independent duplication events.

2. PTOX and the alternative oxidase (AOX) are structurally similar, as expected

from their homologous function, but they display profound differences from

either a biochemical or enzymatic standpoint.

3. The carotenoid biosynthesis hypothesis straightforwardly relates the lack of

PTOX to a defect in chloroplast biogenesis but fails to account for the fact

that some plastids apparently cope with the lack of plastoquinone (PQ)

oxidase, as evidenced by a variegated phenotype.

4. Because chlororespiration controls the redox state of the PQ pool in

darkness, PTOX is a key enzyme in the bioenergetics of the photosynthetic

cell

5. Together with NADH dehydrogenase-like (NDH)/NAD(P)H dehydrogenase-

like (NDA), PTOX sets the redox poise in the dark not only of the PQ pool but

67

also of the entire stroma and thereby potentially impacts, to different degrees,

all the processes that either involve redox reactions or are redox controlled.

DISCLOSURE STATEMENT

The authors are not aware of any affiliations, memberships, funding, or financial

holdings that might be perceived as affecting the objectivity of this review.

ACKOWLEDGMENTS

We thank Benjamin Bailleul and Anja Krieger-Liszkay for critical reading of the

manuscript. W.J.N., F.R., and F.-A.W. acknowledge the support of the CNRS

and Université Pierre et Marie Curie, Paris 6. A.T. acknowledges the support of

CNRS. This work was supported by the Initiative d’Excellence program from the

French State (grant DYNAMO, ANR-11-LABX-0011-01). W.J.N. is a recipient of

a PhD fellowship from Université Pierre et Marie Curie, and N.J.T. acknowledges

support from the Dynamo consortium (Grant DYNAMO, ANR-11-LABX-0011-

01).

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New data about PTOX in the literature

The above manuscript has been published early in 2015. Since then, several studies

further contributed to the knowledge about PTOX and chlororespiration, which gives

me the opportunity to report novelties as detailed below.

A most interesting study came from the group of Krieger-Liszkay (Feilke et al, 2016).

The authors examined tobacco plants with constitutively overexpressed

Chlamydomonas PTOX1 (one of the two paralogs, the one with lower VMAX). A

number of phenotypes are presented, including a significant growth retardation and

loss in chlorophyll content. Most importantly, the authors proposed that PTOX only

attaches to the thylakoid membrane in basic pH that could correspond to the pH of

the stroma upon illumination. In this way, it would contribute to the oxidation of the

PQ pool in high light to alleviate its overreduction, but it would not wastefully oxidize

the pool in light-limiting conditions. This is however not the case for Chlamydomonas,

where a detachment of PTOX from the membrane would result in an inability to ever

detect chlororespiration. It will be also experimentally shown in the section “PTOX in

fluctuating light paper” that PTOX is strongly bound to the membrane in darkness.

These observations are however hard to reconcile with other phenotypes observed in

the overexpressor, most importantly the Phi PSII which should increase if an

additional oxidase is acting as an electron sink (especially because PTOX in these

plants is more abundant than in the WT Chlamydomonas, as normalized to the

chlorophyll content: (Ahmad et al, 2012)), whereas the contrary is observed. Because

the PSI increases simultaneously, increase in CEF is a plausible hypothesis, yet

when the leaves are dark-adapted the oxidation of PSI upon far-red illumination is

faster and not slower as would be expected if CEF indeed increased.

Because the PQ pool is more oxidized in the overexpressor in growth light (when

essayed by HPLC), and a considerable amount of the quinones are sequestered in

plastoglobules instead of being present in the membrane, one should consider the

possibility that the overexpression of PTOX led to a change in the supramolecular

organization of the thylakoid membranes, leading to a disconnection of some PSII

centers from the oxidative activity of PSI. This could readily explain the low PSII,

oxidized pool, low carbon fixation as well as growth and PSI donor side limitation.

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Another recent contribution was provided by the group of Steven Rodermel, who

explored the fate of spots of leaves in PTOX Arabidopsis plants, which mis-

developed as a result of the mutation (Pogorelko et al, 2016). The variegation,

resulting in white leaf sectors in plants grown in moderate and high light intensities is

a consequence of exceeding the threshold of reducing pressure – although it is not

completely clear how this influences plastid development. The authors show that this

development program transforms the plastids from producers to sinks of chemical

energy, and that they still replicate chloroplast DNA, transcribe RNA and express

proteins. These cells from the white sectors have however different cell wall

composition, which increases their susceptibility to bacterial infection. Finally, it is

proposed that the mis-development of plastids is a result of altered retrograde

signaling, possibly by ROS, during the early stages of chloroplast maturation in the

mutant.

Furthermore, a paper from the Grossman lab presents the functional consequences

of nitrogen starvation on the photosynthetic apparatus (Saroussi et al, 2016).

Although our laboratory and others have previously shown that nitrogen deprivation

results in an increase of chlororespiration and a remodeling of the photosynthetic

apparatus in green algae (Peltier & Schmidt, 1991; Wei et al, 2014), the authors

concentrate on the functional aspects of photosynthesis in these conditions.

Interestingly, using mutants of NDA2 and pgrl1 – two of the major actors of CEF –

they show that an increase in P700+ reduction upon nitrogen starvation in the

absence of LEF is solely due to NDA2. They also show that the appearance of qE,

the so-called energy-dependent fluorescence-quenching component, which is related

to the extent of lumen acidification, develops largely thanks to the NDA2 activity in

the light, suggesting a role of an NDA2-mediated CEF despite the low maximal rate

of this enzyme.

One should mention also an effort to elucidate the role of PTOX in non-green

plastids. Recently, a PTOX cDNA has been isolated from carrot roots (Campos et al,

2016), where it was shown to have an expression correlated with genes involved in

carotenoid biosynthesis. A recent review covers this part of research on PTOX which

does not focus on its role in photosynthesis (Renato et al, 2015) and suggests that

PTOX might be the terminal oxidase in a simplified complex I – PTOX – ATP

synthase respiratory chain.

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A very recent publication also describes the involvement of PTOX in the greening of

etioplasts in Arabidopsis. Kambakam and colleagues (2016) report that in the

absence of PTOX the process of chloroplast development from the non green

precursors, etioplasts, is slower, and that PTOX participates in a respiratory activity in

these organelles, which they term the etiorespiration. PTOX seems also responsible

for alterations in chlorophyll biosynthesis, together with the already reported

carotenoids biosynthesis. The former was also observed by Houille-Vernes and co-

authors, who reported that in a double PTOX2/cyt. b6f mutant (Houille-Vernes et al,

2011) cells exposed to even medium light intensities became brown during their

growth on plates.

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Results and discussion

Chlororespiration and its physiological role

Physiological observation

The main goal of the project at the beginning of my internship was to elucidate the

question that was evoked in the introduction: what is the purpose of chlororespiration,

a process that is potentially futile?

We decided to tackle this issue with a physiological approach. We reasoned that, by

exploring multiple growth conditions of the model organism Chlamydomonas, we

could discover instances in which chlororespiration is regulated. Such information is

of course of great value as it can potentially tie an environmental cue to the

relevance of the process. Because it had been already suggested that nitrogen

starvation leads to an upregulation of chlororespiration (Peltier & Schmidt, 1991), one

could assume that also other environmental changes, such as carbon availability,

temperature or light variations, could lead to an increase in chlororespiratory activity.

Similarly, a downregulation of the activity of either of the two light-independent PQ

oxidoreductases – PTOX or NDA2 could bring important indications. If one managed

to obtain several conditions of up- and downregulation of the process, it would pave

the way for a better understanding of its physiological role.

However, already Bennoun had understood the difficulties of measuring

chlororespiration (Bennoun, 2001). Because of its low activity, an order of magnitude

lower than mitorespiration, and because of the cross-talk between mitochondria and

chloroplasts, this task still is difficult to achieve. In our case, changes in the

environmental conditions can likely result in alterations of the cellular metabolism

influencing O2 consumption. Blocking selectively mitochondrial electron transfer is not

a way to address this issue since it strongly modifies the redox state of the

chloroplast (because of the crosstalk between the two organelles, e.g. (Gans &

Rebeille, 1988; Rebeille & Gans, 1988; Rebeille et al, 1988)), increasing the

reduction state of its stroma. This occurs because upon a drop in ATP concentration,

some of the glycolytic enzymes are no longer inhibited allosterically by ATP

molecules, which in turn increases the rate of sugar catabolism, and results in

shuttling of the reducing equivalents to the chloroplast stroma. Alternatively, one

85

could think of working with isolated chloroplasts but not only is this a challenge in the

case of Chlamydomonas, but it also casts doubts on the physiological significance of

such measurements, because of the aforementioned redox cross-talk.

The use of time-resolved spectroscopic approaches in vivo allows one to overcome

these difficulties. It only requires the use of a high cell concentration. Fluorescence

Emission spectroscopy reveals a multitude of information essentially about

photosystem II, yet it also tells about the state of the whole electron transfer chain,

and it provides data about state transitions and the relative lumen pH. Absorption

spectroscopy, especially with the tools developed in our laboratory throughout the

years, allows assessment of the redox state of cyt. b6f, PC and PSI, as well as the

the electric component of the transmembrane electrochemical potential.

Because chlororespiration alters the redox state of plastoquinones, fluorescence

emission spectroscopy is a convenient mean to monitor this process. This is because

PQ pool (Em,7=0mV) is in equilibrium with the QB quinone reversibly bound to PSII,

which in turn equilibrates with the primary electron acceptor in PSII, QA (Em,7=-80mV;

the equilibrium constant of ~20 between PQ and QA). Although the two are in perfect

equilibrium only in the dark, fluorescence is a good proxy for an estimation of the

redox state of the pool. Because most of the chlorophyll fluorescence comes from

photosystem two, and virtually the total extent of the fluorescence variations is a

direct result of changes in the redox state of QA, any alteration of the redox state of

the PQ pool will have an effect on the fluorescence emission. And, following this

logic, when all of the QA quinones are oxidized, the photochemical quenching ability

is the highest and PSII fluorescence exhibits the lowest yield, corresponding to a

basal signal termed F0. On the other hand, when the PQ pool and QA are in their

reduced states, there is no electron acceptor available yielding the so-called ‘closed’

reaction centers. Because of that, the chlorophyll excitation is not converted to

electrochemistry by charge separation (Chl+, Acc-), and the excited state Chl* returns

to the ground state, Chl, by heat or fluorescence. The probability, however, of non-

photochemical and non-radiative excitation loss is thought to be constant in PSII,

therefore a lack of photochemistry results in an increase of fluorescence-mediated

excitation decay. When all the centers are closed, fluorescence emission will attain

its maximal level, FMAX.

86

Moreover, there is a linear relation between an integral of the variable fluorescence

over time and PSII photochemical yield (Bennoun, 1982), allowing quantitative

measurements of the size of the reducible pool of electron acceptors throughout time.

The number can then be related to an actual number of electrons by the mean of

comparison with a DCMU-treated sample, which only allows a transfer of a single

electron before saturating each QA and attaining the maximal fluorescence yield,

FMAX.

However, the experimentalist faces a problem: simultaneous oxidation of the PQH2

by PSI increases the photochemical yield of PSII due to a production of the oxidized

electron acceptors, PQ. Because the CBB cycle then uses CO2 as the terminal

electron acceptor, this PSII yield never decreases and therefore measurements of

the number of electrons needed to reduce the electron acceptor pools would actually

reflect the reduction of every available CO2 molecule. Therefore, one needs an

experimental tool to block the electron transport chain relatively “close” to PSII, in

order to restrain the reducing equivalents to pools of intersystem electron carriers.

In higher plants the above issue is easily solved, as the CBB cycle half-time of

activation is of order of tens of seconds. Thanks to that, there is a drop in PSII

photochemical yield, which approaches the FMAX level of fluorescence upon reduction

of all CBB intermediates upstream the bottleneck, NAD(P)H pool, PSI, PC, b6f and

PQ pool. This feat has allowed Trouillard et al (2012) to establish the kinetics of

chlororespiratory enzymes in a higher plant.

In green algae, the CBB cycle is very rapid to activate - or, as proposed recently (J.

Alric, personal communication) Chlamydomonas similarily to the cyanobacteria

(Allahverdiyeva et al, 2013) express flavor-diiron proteins oxidizing NAD(P)H before

an activation of the CBB cycle occurs - which restrains the experimenter to the use of

mutants. Bennoun (1982) used a PSI mutant for the measurements of the area

above the fluorescence curve that allowed him to observe redox changes of the PQ

pool. Houille-Vernes et al (2011) used a cyt. b6f mutant and a double ptox2/cyt. b6f

mutant in order to quantify the influence of PTOX on the PQ pool and determine its

kinetic properties such a VMAX.

Nevertheless, as stated in the title of this chapter, we aimed at a quantification of

chlororespiration in physiological conditions, where non-autotrophic mutants are of no

87

use. In order to achieve that, we wanted to develop a protocol, which takes

advantage of detectable changes of the redox state of PSI. We reasoned that upon

illumination, P700, the primary donor of PSI, would take longer to become fully

oxidized in the PTOX2 mutant due to an increased quantity of electron donors,

since the PQ pool would be fully reduced in darkness owing to the quasi-absence of

PTOX.

Here, I should state that I will distort the chronology of our discoveries. The results of

the quantification will be described in detail in the next chapter, even if they proved to

be mostly informative about processes other than chlororespiration itself. Luckily

nonetheless, these experiments drew my attention to the fact that the PTOX2

mutant remains in a more reduced state after a minute of darkness, which followed

an illumination, than a wild-type strain. This prompted us to investigate whether this

strain would exhibit growth retardation with regards to the WT in fluctuating light, a

feat never shown for a chlororespiratory mutant. Results of this experiment and our

investigations that followed will be presented here in a form of a pre-publication

manuscript.

88

PTOX in fluctuating light paper

Running titles:

Chlororespiration in Chlamydomonas contributes to the oxidation of PSI

acceptors in the dark and to photoprotection in fluctuating light conditions

Chlororespiration in Chlamydomonas oxidizes PSI acceptors and enables

photosynthesis in fluctuating light

Chlororespiration in Chlamydomonas oxidizes PSI acceptors and enables the

onset of photosynthesis

Photoprotective role of chlororespiration in Chlamydomonas in fluctuating light

Chlororespiration oxidizes chloroplast stroma and contributes to photoprotection in

fluctuating light in Chlamydomonas

Chlororespiration consumes excessive reductants and contributes to photoprotection

in Chlamydomonas

A redox balance between PSI donor and acceptor sides provided by

chlororespiration contributes to photoprotection in fluctuating light

Chlororespiration enables photosynthesis in fluctuating light in Chlamydomonas

89

Authors: Wojciech, Felix, Pierre, Benjamin, Fabrice, Francis-André

1. Introduction

Plants need to cope with large variations in the light irradiance in order to maintain an

efficient photosynthesis. There is a multitude of regulatory processes in the

chloroplast that ensure a maximal possible rate for the linear electron flux from

photosystem II (PSII) to carbon fixation, and at the same time a protection against

over-reduction of the chain, which otherwise would lead to production of deleterious

reactive oxygen species (1-3). These include two prominent types of processes: (i)

downregulating the photochemical activity of PSII, in short term either by quenching

of the excitonic energy (energy-dependent non-photochemical quenching, called qE

(4) or by decreasing its absorption cross-section (state transitions, ST)(5, 6); and (ii),

rerouting the surplus electron flow from PSII to alternative sinks, mostly oxygen

(water-to-water cycles such as Mehler reaction or photorespiration)(7). Cyclic

electron flow (CEF), which reinjects electrons from the reduced ferredoxin back to

plastoquinone (PQ), contributes to both these types of regulation (8, 9).

The aforementioned mechanisms regulate the electron flow in the light, yet there

exist also adjustment processes independent of photosystems’ activities. One of

them is chlororespiration, defined as light-unrelated oxygen consumption in

chloroplasts (10). It is now well established that chlororespiration consists of two

types of enzymatic activities that act in series and share the substrate, reduced or

oxidized PQ and that it is conserved in the photosynthetic lineage. An NADPH:PQ

oxidoreductase (NDA2 in algae, NDH in higher plants) reduces the quinones with

electrons coming from NADPH (11), and a PQH2:O2 oxidoreductase, PTOX (plastid

terminal oxidase) oxidises them (12). Altogether, the in-series activity of NDA2/NDH

and PTOX transfers electrons from the stromal reductants to O2, and their differential

activities regulate the redox state of the PQ pool in the dark (13). In higher plants,

PTOX is implicated in carotenoid biosynthesis during chloroplast development, but its

presence in adult organisms suggests that it continues to play a physiological role

(14, 15). This role is however still not understood.

Chlororespiration, intertwined with the electron transfer chain by sharing PQ, is an

antagonist of the light reactions of the photosynthesis, consuming its primary

products, O2 and NADPH (12). The maximal rates of chlororespiratory enzymes are,

90

however - depending on the species - at least two orders of magnitude lower per

chain unit than the rate of the slowest step of electron transfer, oxidation of a PQH2 at

the Qo site of cytochrome b6f (cyt. b6f) (13, 16). The low rates of PTOX and its

counterpart PQ reductase suggest that they can hardly compete with the other PQ

reductase (PSII) or reductase (cyt. b6f) in the light, and therefore that the role of

chlororespiration in the light is limited. However, previous studies showed that a

mutant of the major chlororespiratory oxidase in the green alga Chlamydomonas,

PTOX2, exhibits no difference in fitness when grown in the dark even though its

PQ pool remains reduced (13). The relation between chlororespiration and other

regulatory mechanisms is also poorly understood. Here, we show that PTOX2,

whose in vivo rate per photosystem is at least 10x higher than in higher plants, does

not play a significant role during steady-state illumination, yet its activity is crucial

under fluctuating light. Chlororespiratory activity oxidizing the stromal reductants in

darkness makes the onset of photosynthesis faster and more efficient, and strongly

influences regulatory mechanisms such as CEF.

91

2. Results

PTOX2 does not significantly contribute to ETR in the light

One can expect benefits from quinol-oxidating activity in conditions where the

capacity of the chloroplast to reduce carbon (IV) is lower than PSII light absorption,

which prompted earlier reports to suggest a photoprotective role of PTOX (14, 17,

18). Because our group has shown before that the rate of PTOX is incompatible with

an electron sink role (13), we first sought to reinvestigate these claims by monitoring

the electron transfer rate (ETR) in the WT and

intensities. Indeed, if PTOX accepts a significant proportion of the electron flux from

PSII, then the maximal ETR should be significantly higher in the wt than in the

mutant. As shown in Fig. 1, the presence of PTOX2, the major PQ:O2 oxidoreductase

in Chlamydomonas does not significantly contribute to the ETR in the steady-state at

any light intensity in cells cultured in TAP.

Figure 1. Electron transter rate in WT and DPTOX2 over a range of light intensities. ETR is

calculated from the PSII parameter after 30s adaptation to each light intensity. See Materials

and Methods section for details.

92

PTOX2 mutant exhibits severe growth phenotype in fluctuating light

Photosynthetic microorganisms are subjected to irradiance conditions which are

more severe that those of higher plants. Because PTOX2 mutant does not exhibit a

growth phenotype in darkness despite its lock in a state II, and because of its low

maximal rate we investigated growth of the mutant in fluctuating light conditions. As

shown in the Fig. 2, growth rate of PTOX2 is severely decreased in such conditions

with regards to the WT in a medium supplied with reduced carbon, TAP. Importantly,

the growth of the mutant is not compromised in minimum medium in fluctuating light,

nor in continuous illumination, nor in darkness (Fig. S1). Together, these results

highlight a crucial role of chlororespiration in darkness and during the dark-to-light

transitions and also argue against significant PTOX role in continuous illumination.

Figure 2. Growth of the WT and DPTOX2 in fluctuating light conditions. A, a representative

growth curve of the WT and the PTOX mutant. B, growth rate of the two strains in fluctuating

light (n=2± S.E.).

93

These results led us to explore the contribution of chlororespiration during dark/light

transition periods. We started by examining the distribution of PTOX in the

chloroplasts - according to a recent report, PTOX in higher plants in the dark

(supposedly lower stromal pH) would detach from the thylakoids and become stroma-

soluble, therefore it would be unable to oxidize quinols. As shown in Figure 3, in both

light and darkness, we found the protein in the membrane fraction of the chloroplasts,

in contrast with previous observations in plants (19).

Figure 3. Fractionation of whole cell protein extracts isolated in the light and in darkness.

PTOX2 – expectedly for a peripheral membrane protein - is detected in the total and membrane

fractions of Chlamydomonas extracts in both light and dark conditions, similarily to cyt. b6f

subunit IV, a transmembrane protein. Thioredoxin H is a soluble protein and it is not detectable

in the membrane fraction.

94

Because neither continuous darkness nor illumination and only fluctuations prove to

be detrimental to a chlororespiratory mutant, we investigated the influence of PTOX

during both dark- and light periods using functional approaches. We developed a

treatment inside the spectrophotometer that mimics the conditions of fluctuation

where PTOX2 exhibited growth differences with regards to the WT, and we

measured photosynthetic parameters throughout this regime, until a steady-state

level was achieved.

Given the role of the redox state of the PQ pool in the regulation of state transitions

(13), WT and mutant cells were first adapted to low light intensity, setting both of

them in state I prior to the fluctuating light treatment (Fig. S2a), a feat not achievable

in darkness.

QA redox state and P700 oxidation demonstrates stromal reductants accumulation

during fluctuations in the absence of chlororespiration

We started by investigating the redox relaxation of the photosynthetic electron

transfer chain during the dark periods and first explored the rate of QA reoxidation

during the first and last dark period of the 30 min. treatment. We have applied the

Stern-Volmer relationship to quantify the concentration of QA- from the fluorescence

data (see supplementary materials for the equations) after applying a saturating

pulse to reduce 100% of the QA quinones. As shown in the Figure 4A, QA, which is

fully reducedafter the saturating pulse, is nearly completely (WT) or partially oxidized

after the 1 minute darkness period (PTOX2). That is because quinol oxidizing

activity has a higher rate than the NAD(P)H:PQ oxidoreductase. However, in the

absence of PTOX2, the oxidation of the PQ pool is much less efficient and 40% of

PSII centers – in equilibrium with the PQ pool - are in their reduced state at the end

of the dark period. The biphasism of the QA oxidation is expected (see discussion for

more details) and the slower phase corresponds to the PTOX-mediated PQ

oxidation.

The kinetics of the QA reoxidation is slower after the 30 min. light/dark treatment,

suggesting an accumulation of NADPH in such regime, and QA is in all instances

more reduced in the mutant (Fig. 4B).

95

In order to confirm the hypothesis that the NADPH accumulates in the stroma in

fluctuating light, and that it is more pronounced in PTOX2 due to the decreased

chlororespiration we have measured the acceptor side limitation of PSI at the end of

the dark period throughout the 30-min treatment using the Klughammer and

Schrieber protocol (20).

Figure 4. The kinetics of QA- concentration changes in darkness after a saturating pulse in

fluctuating light as determined from the fluorescence decay kinetics. The maximal quantity of

quencher (corresponding to the [QA-] value of 0) reflects the low-light adapted condition

followed by a brief dark-adaptation. A, after the first illumination period; B, after 30 minutes of

fluctuating light treatment.

96

The results are shown in the figure 5. As predicted by the hypothesis of an

accumulation of reducing equivalents, during the treatment the quantity of

photooxidizable P700 decreased in both strains. However, in the PTOX2 the

decrease is sharper and to a bigger extent than in the WT, evidencing the necessity

of chlororespiration in the maintenance of a redox homeostasis of the stroma before

an illumination.

All the measurements so far pinpoint to a crucial role of chlororespiration in the dark

periods, which allow to reoxidize QA (Fig. 4), PQ pool and, in turn, PSI acceptors (Fig.

5) prior to the illumination. Can these results explain the decrease in growth rate in

the fluctuating light in the mutant compared to the wild-type? In a case where at the

onset of illumination, PSI centers are closed because of a stable overaccumulation of

their electron acceptors, corresponding PSIIs would also be blocked, thus decreasing

details).

Figure 5. Effect of the fluctuating light treatment on the quantity of photooxidizable P700. The

maximal quantity of P700+ probed during a 20ms saturating pulse was normalised to a DCMU-

treated reference. See text for details.

97

Fluctuating light and excess of reductants compromise PSII activity in the PTOX

mutant

As a mean to measure the redox changes in the electron transfer chain we have

followed the changes in some fluorescence parameters during the fluctuating light, as

shown in figure 6A.

The three parameters acquired during the fluctuating light treatment are (i) FM

(normalized to its initial value, see Methods) (ii) FV/FM at the beginning of each light

period; (iii) PSII at the end of each light period.

As shown on the Fig. 6B/C, 60s light/60s dark treatment for 30 minutes induces

significant changes in the fluorescence parameters. The FM decrease is minor and

nearly identical in both strains. There are at least two possibilities for this drop: (i)

photoinhibition, which yields low-fluorescence PSII centers and (ii) state I to state II

transition, where the PSII antenna, LHCII, migrate to non-appressed regions of the

thylakoids and increase the PSI cross-section. We decided to decipher between

these two hypotheses by two means: we have measured the 77K fluorescence

emission spectra before and after the treatment which show that both strains undergo

a state I to state II transition throughout the treatment (Fig. SX), and, consistently with

the FM data, the extent of the transition is slightly higher in the WT than in the mutant.

Because 77K fluorescence spectra are not quantitative, we have also measured the

FM relaxation after the treatment in both strains. In darkness, the FM slowly recovers

in the WT, whereas in the PTOX2 it decreases even more than during the

treatment, consistent with the mutant being locked in state II (data not shown).

However, when a low intensity far-red light was applied, preferentially exciting PSI,

the PQ pool should get oxidized regardless of the presence of PTOX. Indeed, in both

strains the FM increases rapidly (within ~3 minutes), eventually recovering to the pre-

treatment level. This proves that virtually all of the change in FM during the treatment

is due to state transitions.

The value of FV/FM before the treatment is unusually high for Chlamydomonas (0.82),

and higher than in a dark-adapted sample. Both strains are in perfect state I as

evidenced by the 77K fluorescence spectra before the treatment, and as FV/FM does

not change in PTOX2 following a brief dark-adaptation. Those observations suggest

that the samples are lacking reducing power in the chloroplast after the low-light

98

adaptation. Such pre-treatment is optimal as for both WT and PTOX2 the starting

point is identical, at odds with previous comparisons of those two strains using a dark

adaptation pretreatment (13).

During the fluctuating light treatment, the FV/FM only slightly decreases (~5%) in the

WT but there is a decrease of about 30% in PTOX2. This is consistent with the

redox state of QA at the end of the dark treatment (Fig 4).

Surprisingly, the PSII parameter, which reflects the PSII photosynthetic electron

flow (21) follows the same kinetics as the FV/FM variations. The photochemical yield

in the PTOX2 mutant diminishes by around 40%, whereas no significant change was

observed in the WT. This indicates a crucial role of PTOX – or chlororespiration -

under these fluctuating light conditions and readily explains the differences in growth

rate between the two strains.

In order to test whether the frequency of the fluctuating light treatment influences the

phenotype described above, we have decided to investigate 1s/1s dark/light cycles

which maintains the maximal light intensity and the total photon quantity. As shown in

the Figure 6C, this regime does not lead to any differences in FV/FM or PSII

between the two strains during the treatment. This result proves that in order to

reveal the influence of chlororespiration, a longer period of darkness is necessary,

which is in agreement with the low rate of the process. As shown in the figure S4,

cycles of 20s/20s and 40s/40s are already sufficient to manifest the maintenance of

the redox state of the electron transfer chain and the stroma by chlororespiration,

once again consistent with the rate of chlororespiration as shown in Fig. 4.

In order to investigate the influence of the overall redox state of the chloroplast, we

have repeated the 60s/60s light treatment with cells grown in minimum medium, i.e.

without an addition of a reduced carbon source. The differences between the two

strains throughout the treatment were less important than in the TAP-grown strains,

important when the chloroplast deals with an important flux of reducing power coming

from the cytosol. As this treatment did not induce significant changes in growth rate,

we conclude that chlororespiration essentially alleviates the reducing pressure in the

chloroplast stroma by discarding excess electrons.

99

However, none of our observations can explain the decrease in PSII in the PTOX2

mutant. In light, the redox state of PQ pool is determined by the rates of reduction

and oxidation by PSII and cyt. b6f, respectively; similarily, NAD(P)H pool state is

determined by its reduction by PSI and oxidation essentially by the CBB cycle, not

chlororespiration, so the initial redox states of the two cannot account for the PSII

decrease.

One possibility for the PSII difference between the WT and the PTOX mutant would

be however that PTOX activity increases significantly during the course of the

fluctuating light treatment, allowing an electron sink role in such regime. If this is the

case, the rerouting of a significant proportion of the electron flux from PSII should

decrease by as much the PSI flux.

100

Figure 6. Measurements of fluorescence parameters allow the determination of the PSII status

and activity in Chlamydomonas cells in fluctuating light. A, fluorescence trace during a light

period. Black bar, dark period; yellow bar, actinic light; orange bars, saturating light pulse

(~5000uE.m-2.s-1). This protocol was repeated at each cycle and the fluorescence trace was

used to calculate the fluorescence parameters presented in panels B and C. Fluorescence

parameters throughout 60s/60s (B) or 1s/1s (C) 0/340uE fluctuating light treatment.

101

The overreduction in PTOX results in an establishment of CEF-favourable

conditions, resulting in an increase in PSI

Contrary to the measurements of P700 at the beginning of the illumination, PSI

acceptor side becomes more oxidized in the PTOX2 mutant at the end of the light

period (Figure 6). Indeed, throughout the treatment: the quantity of photooxidable

P700 increases with regards to the WT, and there is an accumulation of oxidized

P700 in the light.

Figure 7. PSI parameters during the fluctuating light treatment. Absorption at 705 nm was

monitored during the light-to-dark transitions in the WT and in the PTOX2 mutant. Fraction of

photooxidizable P700 was determined using a short saturating pulse at the end of the light

period; P700+ accumulation in the light corresponds to the difference between absorption

before the pulse and in darkness. Both quantities are shown as a fraction of total P700 as

determined in the DCMU-treated sample in the light with a saturating pulse superposed.

102

We have then examined the PSI at the end of the light period. It corresponds to the

fraction of “open” PSI centers, i.e. P700 which is reduced in the light but is ready to

perform photochemistry (photooxidable under a saturating pulse), and is therefore

proportional to the electron flux through PSI. Surprisingly, PSI parameter also

increases during the treatment, and the increase is far more pronounced in the

mutant than in the WT. The exact opposite was observed for PSII (Fig. 3). These

results disprove our initial hypothesis that PTOX is responsible for the higher PSII

photochemical yield in fluctuating light in the WT compared to the mutant.

As the parameter provides only the fraction of photosystems that are performing

photochemistry, it cannot be used to directly compare the electron transfer rates

flowing through respective centres. The electron flow should be calculated as the

product of the photochemical yield by the light absorption by both PS, and therefore

requires the measurements of the absorption cross-sections of PSI and PSII (see

equation in the supplementary material). Although we try to estimate those (see

supplementary discussion), any adjustments due to state transitions, which are the

reason for the Fmax decrease during the treatment, will only increase the differences

between PSI and PSII fluxes because upon ST, PSI antenna increases at the

expense of PSII antenna size.

To reconcile the observations that in the PTOX2 mutant during the treatment (i)

PSII decreases, and (ii) PSI increases, one needs to evoke a pathway that

requires PSI photochemistry but not PSII. Cyclic electron flow is the obvious

candidate – and it has been suggested that simultaneous measurement of PSII and

PSI is the most correct way to assess CEF after antenna size adjustment and ETR

calculation (22). We can estimate the CEF rate by comparing the respective PSI and

PSII fluxes, knowing that PSII contributes exclusively to LEF and the PSI to

LEF+CEF in the light. In the case of WT, CEF in the steady-state of the treatment

contributes little to the PSI flux, on par with estimates from continuous illumination in

the algae – the (PSI-PSII)/PSI ratio is very low (ref). In PTOX2, however, its

contribution to PSI flux is much more significant as more than a half of the electrons

transferred by PSI are of CEF origin. Because the parameters are not adjusted to

the antenna sizes, the absolute value of CEF rate can only be underestimated (see

more in supplementary discussion). Finally, the high CEF rates are consistent with

103

our obserations that the PSI acceptor side is oxidized at the end of the light period

and by the accumulation of P700+.

Interestingly, although it was reported that PSI can be damaged when exposed to

fluctuations in light and/or acceptor side limitations, we were not able to see any

significant photoinhibition of PSI during the treatments (data not shown). We

hypothesize that this is due to an insufficient light quantity received, since light is

applied only for 50% of the duration of the treatment.

WT DPTOX2

Beginning of

the treatment

210 204

End of the

treatment

224 176

Table 1. Influence of the treatment on the photochemical rates (expressed in e-.s-

1.2PS-1) in WT and DPTOX2.

and PSI were measured at the end of the 1 min light period (n=3-4)

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3. Discussion

PTOX in Chlamydomonas cannot compete for electrons with PSI

PTOX activity can be seen two-way. In darkness, although oxidizing directly

only quinols, it is involved in chlororespiration with its counterpart, NAD(P)H

dehydrogenase (23, 24). NDH or NDA2 reduce quinones using electrons from

the chloroplast stroma, functionally forming an NAD(P)H:O2 oxidoreductive

pathway. The lack of PTOX in darkness results in an overreduction of PQ

pool, leading to a lock in state II (13). PTOX is however also active in the light,

which raised proposals about its involvement in photoprotection under high

light REFS. However, in the light it faces competition for electrons of the cyt.

b6f and importantly, the maximal rates of quinol oxidation per electron transfer

chain unit of these two are markedly different (~150 for cyt. b6f to ~5 for PTOX

in Chlamydomonas (12)). We have shown that in any light intensity, PTOX

cannot efficiently compete with cyt. b6f and does not significantly contribute to

PSII ETR, let alone regulate the PQ pool redox state. These observations are

in agreement with previous measurements when upon a saturating flash, due

to a positive charge formation by the PSI, an oxygen consuming pathway in

the chloroplast decreased (25). It must be however kept in mind that it is not

always the case – marine algae and cyanobacteria increase PTOX activity

which contributes significantly to the PSII flux (26, 27).

In Chlamydomonas, the role of chlororespiration is to consume excess

reductants in the dark

A lack of significant activity in the light and no effect on growth in the PTOX2

mutant in darkness (13) strongly points to the importance of this protein in a

fluctuating light environment. We have shown that apart from a strong effect

on primary photochemistry in the mutant, as discussed before, PTOX2

exhibits decreased growth rates with regards to the WT in fluctuating light.

PTOX-driven PQ oxidation joins here a number of other regulatory

mechanisms reported recently (28), which prove useful beyond standard

laboratory conditions. Flavodiiron proteins seem in particular to perform a

similar function in cyanobacteria, where they serve as a typical overflow

105

device – oxidizing part of NAD(P)H when a saturating light is superposed on

growth light (29). Similarily, state transitions-mediated regulation of electron

flux is important for the growth in fluctuations or overreducing conditions (30,

31). Closer to PTOX’s function, in cyanobacteria multiple mutants of terminal

oxidases exhibit growth phenotype when exposed to fluctuating light when the

intensity is high (32), but photosynthetic parameters were not measured for

these cultures. Similarily, photosynthesis in fluctuating light conditions in the

CEF mutants is impaired (33) – the authors showing that PSI is the target of

photoinhibition in such case. In our experience, total PSI quantity never

decreased despite PSI acceptor side overreduction upon the illuminations.

In our experimental conditions we observe a decrease in the FV/FM value in

the PTOX2 mutant during the treatment, which we interpret as an

accumulation of reductants in the chloroplast stroma, which in turn influences

the redox state of QA quinone in PSII. As discussed before, the PQ and

NAD(P)H pools are connected by NDA2. We relied on an analysis of the

multiphasic decay of fluorescence after a saturating pulse to determine the

rates of quinone oxidation before- and in the steady-state of the treatment

thanks to a conversion of the fluorescence data to [QA]. After a first, rapid

phase of oxidation of small amplitude, which we interpret as a reduction of

reduced QA and PQ thanks to the activity of the cyt. b6f, a longer phase of

greater amplitude occurs. The latter is an oxidation of the remaining quinols

mediated by PTOX. We assume that the oxidation rate did not change

throughout the treatment (although PTOX was proposed to produce ROS

(krieger_liszkay) and its enzymatic cycle involves radical compounds (moore)

and interpret the increase in QA- concentration as an increase of NDA2 rate

due to an increased concentration of its substrate, NAD(P)H.

It is interesting to hypothesize about the kinetics of the establishment of the

steady state FV/FM value during the fluctuating light treatment. This process is

long and takes approx. 15 minutes, or approx. 8 cycles of illumination in the

PTOX mutant. Taking into account the light intensity, it corresponds to tens of

thousands of PSI photochemical events before achieving a redox steady-

state. Although many of these electrons are consumed by the CBB cycle,

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such a vast number is incompatible with the size of NAD(P)H pools, therefore

we conclude that the electrons influencing the redox state of the stroma can

be at least partially stored in compounds in equilibrium with NAD(P)H,

presumably the metabolites of CBB.

Photosynthetic electron flux is drastically decreased in a PTOX mutant in

fluctuating light

Using a set of in vivo biophysical approaches and growth phenotyping we

have determined that although the overall photochemical rate is only slightly

lower in PTOX2, the PSII-driven electron transfer is strongly decreased

which in turn impedes growth of the algae. One can formulate several

hypotheses for this phenotype: (i) PTOX2 plays a significant quinol oxidase

role in the light; (ii) PSII photoinhibition decreases PSII activity; (iii) part of PSI

compartments being disconnected from oxidation by Calvin-Benson-Bassham

cycle, resulting in a block of some chains; and (iv) long-term ultrastructural

changes in the physiology of photosynthetic electron transfer mediated by the

stromal redox state. The first hypothesis fails to account for the fact that the

maximal rate of PTOX is incompatible with significant differences in QA

oxidation state; PSII photoinhibition is also not the case in our conditions as

the FV/FM decrease throughout the treatment is reversible in timescales

corresponding to state transitions. The third hypothesis is on the other hand

conceivable, yet not without further suppositions. In a homogenous system,

both NADPH and PQ pool upon illumination are oxidized by the PSI and CBB

cycle, respectively, “losing” the memory of their respective initial states on the

onset of illumination. Nevertheless, there exist a lot of structural data showing

that neither the thylakoid membranes, nor the stroma of the chloroplast of

Chlamydomonas are homogenous (Engel eLife). Moreover, although the

exact concentration of PTOX is not known for Chlamydomonas, it was shown

that it is highly sub-stoichiometric in plants with regards to PSII – and even if

the differences in the rates of PQ oxidation between algae and higher plants

are a simple result of PTOX being more abundant in the former, it would still

be less concentrated than PSII.

107

Nonetheless, even if the NADPH, reduced in the compartments inaccessible

to chlororespiration, could remain reduced during the light period and block

entire units of electron transfer chains - as a result, lowering PSII, such

hypothesis is easily tested. We have used P700 oxidation measurements to

determine which fraction of PSI was photooxidable in the light at the end of

the minute of illumination, together with determining accumulation of P700+

accumulated in the light and PSI. The results, shown in fig. 7 contradict this

hypothesis, as every parameter related to PSI redox status appears to be in a

more oxidized state in the mutant at the end of illumination.

Chloroplast overreduction sets it in a cyclic electron flow-favourable mode

The increase of CEF in the PTOX2 mutant in fluctuating light treatment is an

important observation on many levels. First of all, it likely proves that in

physiologically relevant conditions (i.e. without DCMU) the redox state of the

chloroplast is a signal for the increase of CEF. This goes in line with

observations in anoxia (34) where CEF rates increase to contribute to a net

ATP production and as a result, more efficient consumption of NADPH by

CBB cycle. We document chlororespiration as a crucial regulator of the

chloroplast redox maintenance in the dark conditions, not only for the PQ pool

but also for the PSI stromal acceptors, a feature which was never

demonstrated before. It is however difficult to pinpoint the exact source of

signalisation which increases the activity of CEF: in the PTOX2 mutant, not

only the NAD(P)H, but also PQ pool is reduced, similarly to anoxia. Even

though CEF is a PQ-reducing activity – so in short term its effect is inverse of

the sought – it actually helps oxidizing the PQ simply by increasing the rate at

which NADPH is consumed meaning that both of these pools could be a

signal to increase CEF. Joliot & Joliot (35) also hypothesized that ATP/ADP

ratio could be the sensor for CEF regulation.

In a homogenous system, where the PQ pool is common to both linear and

cyclic electron flows, one does not expect neither a Fd:PQ oxidoreductase nor

an NAD(P)H:PQ oxidoreductase to kinetically compete with PSII activity.

Perhaps the most interesting observation is however that CEF is indeed able

108

to compete with LEF in our experiments, supporting the hypothesis that the

two modes do not share the same pool of PQ.

This is substantiated by the observations that although after establishment of

a steady-state of the treatment (after 15 mi

PSII is stable during the illumination, but the quantity of photooxidable PSI

changes from ~0.3 to ~0.8. Because CEF is active during this period, as

evidenced by the PSII/PSI ratio and because the light we used for the

treatment results in a significant fraction of PQ pool reduction, it is concluded

that in order for this efficient competition CEF needs a PQ pool physically

separated from this utilized by PSII. Because no standalone oxidoreductase

which mediates CEF as a Ferredoxin:plastoquinone oxidoreductase has been

found despite long-lasting efforts and because of its high rates (as discussed

in Supplementary discussion) we tend to believe that CEF is mediated as

initially proposed by Mitchell (36-38), via the heme ci in cyt. b6f (39). This

assumption is relevant even if the PQ pool is highly reduced, because one

can envisage that the plastoquinone just oxidized at the Qo site can be used

by site Qi for reduction without diffusing to the PQ pool out of the cyt. b6f

dimer, similarly as it possibly happens in anoxia (Joliot). Such mechanism

would allow for a fast CEF with rates of the same order of magnitude as linear

electron flow, observed in higher plants and algae in anoxia (despite the PQ

pool redox imbalance pointed out by Allen (40)) and would account for the

observation that PQ pool in our experimental conditions is reduced despite

oxidized PSI acceptor side and very rapid CEF.

Otherwise, as suggested before (41-43), because of thylakoid membrane

heterogeneity and photosynthetic complexes’ compartimentation the diffusion

of PQ is locally restricted. Most of the PSI centers and roughly half of the cyt.

b6f complexes are located in the non-appressed regions of thylakoids (44),

which increases the probability that an oxidized quinone despite having

diffused out of the b6f dimer is available for an FQR-mediated reduction.

Regardless of which of those two possibilities accounts for the fact that CEF

efficiently competes with linear flow in our experiments, they cannot explain

the increase of the CEF rate throughout the fluctuating light treatment.

109

Although, as discussed before, it is likely that the accumulation of reducing

equivalents (as evidenced by the FV/FM changes) throughout the treatment

results in increased cyclic, it is inconceivable that a passive reducing pressure

from the stroma, increasing the probability of low-potential electron carriers

reducing back the membrane transporters, can account for that. The reason

for this is the following: we have shown that at the end of illumination in the

PTOX2 mutant there is an accumulation of P700+ and a large fraction of PSI is

photooxidable, yet it is the condition where the CEF is the most prominient.

This strongly suggests that, in line with earlier propositions (42, 45), some

degree of structuration of CEF components is necessary, as,

thermodynamically, the situation most favourable for CEF would be (i) PSI

acceptors reduced, and (ii) PQ pool oxidized.

If indeed the availability of oxidized quinones is not a limiting aspect for CEF,

it is evident that the regulation takes place at the PSI acceptor side, and

determines the probability with which the electrons return to quinones. Two

major types of CEF structuration were proposed in the literature, either

involving a differential binding of FNR to the cyt. b6f depending on the stromal

redox state, or a closing in between PSI and cyt. b6f.

Our experiments in fluctuating light substantiate hypotheses that

chlororespiration is a process regulating the redox state of the chloroplasts

stroma. This may explain the complex evolutionary history of PTOX enzymes

that keep being duplicated, especially in microorganisms (12, 46), often

exposed to fluctuation in light and carbon availability (47), or inhabiting dense

populations with oxygen limitations (where PTOX could act as an oxygen

sensor). It is equally interesting to hypothesize about the rates of

chlororespiration – why it is an order of magnitude lower in higher plants than

in algae (16), contrary to cyanobacteria, which possess additional terminal

oxidases (32)? Plants have retained the complex I-like NDH as a PQ

reductase, and it has been proposed that this enzyme is a Fd:PQ and not a

NAD(P)H:PQ oxidoreductase (48) - the scarce amounts of its substrate in the

dark result in a low reductive pressure on the pool and may explain the low

abundance and/or rates of PTOX.

110

Morever, PTOX2 mutant in fluctuating light proves to be also a perfect

system to study the regulation of CEF, a feat which was hampered until now

by the employment of anoxic conditions, in which case the linear electron flow

from water to CO2 is impeded.

Fig. 9. A model of the situation taking place in fluctuating light in the WT and

in PTOX mutant.

111

4. Materials and methods - unfinished

P700 oxidation

…..To achieve that, we have applied a saturating pulse (therefore avoiding

any variation owing to the antenna size differences) at the beginning of each

light period and detected the redox state of P700 throughout the pulse. We

have then taken the minimal value of absorption corresponding to the

maximal level of oxidation and normalized it to the maximal extent of P700

oxidation (saturating pulse in the presence of DCMU).

We have chosen to follow the light regime which yielded growth differences:

darkness - where chlororespiration is an exclusive mechanism setting the

redox poise of the electron transfer chain, and is likely to have the biggest

impact on subsequent photosynthesis - to medium-intensity, non-saturating

light, in which the PTOX-mediated quinol oxidation is supposed to be

insignificant (Fig. 1). We have used equal periods of light and darkness and

changed the length of a cycle (from 1s of light/1s of darkness, to 180s/180s)

to assure the same average irradiance in all treatments. This kind of light

fluctuations aims at mimicking the most extreme natural variations possibly

encountered by green algae. We have performed the measurements until a

steady-state of the parameters was established

A following protocol has been employed: the cells grown in their respective

medium were concentrated and adapted to low light intensity, setting both the

mutant and the WT in state I (Figure S3a-77K spectra). The cell were briefly

dark-adapted and transferred to the JTS. During the fluctuating light

treatment, each dark-to-light and light-to-dark transition was monitored as

depicted in the Figure 3A: first, the level of fluorescence in darkness was

monitored, yielding the F0; next, an actinic light (AL) of a given intensity was

turned on, and a sa -2.s-1) were superposed to

measure the maximal fluorescence yield Fmax; fluorescence was then

measured until a quasi steady-state was achieved (~5s). The AL was kept on

for the indicated time, at which end the stationary level of fluorescence in the

light was measured (Fs), followed by a saturating pulse (Fmax). The AL was

112

turned off instantly after the saturating pulse, and the fluorescence decay was

measured for 15 seconds.

113

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1. Supplementary data

𝐹𝑣/𝐹𝑚 =𝐹𝑚𝑎𝑥−𝐹0

𝐹𝑚𝑎𝑥 Eq. 1

𝜙𝑃𝑆𝐼𝐼 =𝐹𝑚𝑎𝑥−𝐹𝑠

𝐹𝑚𝑎𝑥 Eq. 2

𝜙𝑃𝑆𝐼 =𝑃700𝑙𝑖𝑔ℎ𝑡+𝑝𝑢𝑙𝑠𝑒−𝑃700𝑙𝑖𝑔ℎ𝑡

𝑃700𝑚𝑎𝑥 Eq. 3,

where 𝑃700𝑚𝑎𝑥 is the value upon saturating pulse in a DCMU-treated sample.

Note that the 𝑃700𝑙𝑖𝑔ℎ𝑡+𝑝𝑢𝑙𝑠𝑒 value is lower due to acceptor side limitation of

P700.

𝐸𝑇𝑅 = 𝜎 ∙ 𝐼 ∙ 𝜙𝑃𝑆𝐼𝐼 Eq. 4,

where 𝜎 is the cross-section of PSII and 𝐼 the light intensity.

According to Stern-Volmer relationship, quencher Q concentration influences

the rate of fluorescence in a following manner:

𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑎𝑏𝑠𝑒𝑛𝑡)

𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑝𝑟𝑒𝑠𝑒𝑛𝑡)= 1 + 𝑘𝑆𝑉[𝑄]

where 𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑎𝑏𝑠𝑒𝑛𝑡) is the fluorescence intensity without a quencher,

𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑝𝑟𝑒𝑠𝑒𝑛𝑡) with the quencher, 𝑘𝑆𝑉 is the quenching coefficient and [𝑄] the

concentration of a quencher.

A transformation of this equation gives:

𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑎𝑏𝑠𝑒𝑛𝑡) − 𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑝𝑟𝑒𝑠𝑒𝑛𝑡)

𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑝𝑟𝑒𝑠𝑒𝑛𝑡)= 𝑘𝑆𝑉[𝑄]

One can substitute 𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑎𝑏𝑠𝑒𝑛𝑡) to 𝐹𝑚𝑎𝑥 (the maximal level of fluorescence

when the QA is in its reduced state after a saturating pulse), 𝐼𝑓𝑙𝑢𝑜 (𝑄 𝑝𝑟𝑒𝑠𝑒𝑛𝑡) to

𝐹, and the [𝑄] to [𝑄𝑎 (𝑜𝑥)], which yields:

𝑘𝑆𝑉[𝑄𝑎 (𝑜𝑥)] =𝐹𝑚𝑎𝑥−𝐹

𝐹, or

𝑘𝑆𝑉[𝑄𝑎 (𝑟𝑒𝑑)] = 1 −𝐹𝑚𝑎𝑥 − 𝐹

𝐹

119

As shown before, a transition from a quasi-perfect state I (PSI:PSII antenna

size ratio of 1:1) to state II occurs during the treatment. As the Fmax changes

are complementary to the increase of PSI antenna size, we roughly estimate

that for the shown 20% PSII antenna size decrease, there is a 20% increase

in the PSI cross-section. The final ratio is therefore 1.2:0.8, (PSI antenna 50%

0.3

These relative contributions to the photochemistry may be then used to

precisely describe the ETR of each photosystem: knowing that the

photochemical rate in the WT is 220e/s/2PS and 170e/s/2PS in the DPTOX2

at the end of the treatment: PSII ETR is 88e/s, and PSI 132e/s (CEF of 44e/s)

in WT, and PSII ETR is 43e/s and PSI ETR is 128e/s (CEF of 85e/s).

120

Trials of quantitative in vivo measurements of chlororespiration

reveal information about CEF in Chlamydomonas

Initial idea overview – measurements of chlororespiratory fluxes

As mentioned in the introduction to the „Results“ section, the main goal of the

thesis was to establish a role of chlororespiration in the physiology of

Chlamydomonas, using the PTOX2 mutant available in our laboratory at the

start of my project. This unicellular algal species is a model organism for

photosynthesis studies due to a unique chloroplast, which bears

photosynthetic apparatus largely identical with this of higher plants. We aimed

at achieving this goal using a functional approach, i.e. by measuring

chlororespiratory fluxes in different conditions, reasoning that correlating

increased chlororespiratory activity with changes in the physiological state of

the cells can provide clues with regards to the relevance of this process.

There are two possibilities of the assessment of the redox state of PQ pool.

Either we can count how many more electrons we can add to completely

reduce the pool - using fluorescence – or oxidize the pool and count how

many electrons were present at the beginning of the experiment using a PSI-

based approach. In both cases, we need to ensure that no leaks are present:

in the first case, leaks that would oxidize the pool during our measurement,

and in the second case, no reducing pathways increasing the reduction.

Measurement of the redox state of PSI was the method of choice due to the

reasons outlined in the introduction. Briefly, fluorescence-based

measurements of the redox state of the PQ pool in Chlamydomonas are not

achievable in the WT. The reason for this is that, in contrast with plant

chloroplasts that exhibit a block in photosynthetic electron flow upon a dark to

light transition, due to the laps of time required for activation of carbon fixation

through the CCB cycle, photosynthetic electron flow develops at high rates in

the same conditions in Chlamydomonas. This is due either to a more rapid

activation of CBB cycle in algae or, alternatively to the presence of flavo-di-

iron proteins as transitory electron acceptors of PSI in the chloroplast stroma

(J. Alric, in preparation). Because we wanted to work in physiological

conditions, the use of photosynthetic mutants – which would block the transfer

121

of electrons to downstream acceptors in the vicinity of PSII - was excluded,

and inhibitors of electron transfer chain such as tridecyl-stigmatellin (cyt. b6f),

iodoacetate, glyceraldehyde (both CBB cycle) etc. were neither reliable, nor

specific enough.

Thus, we decided to employ measurements of the redox state of P700, the

primary chlorophyll donor to PSI, in order to quantify the redox state of the PQ

pool and to gain kinetic information about the rates of its reduction and

oxidation. The redox changes of P700 result in relatively important signals

detectable at 705 nm, which are easily measured, using the absorption

difference spectrophotometer available in the laboratory (JTS-10, Biologic), in

vivo with minimal preparation of the sample. The high potential of the oxidized

special pair in the reaction center, P700+, results in its complete reduction in

darkness (even in the absence of its electron donor, plastocyanin), providing a

reliable starting point for the measurements.

On the other hand, PSI is supplied with electrons by the photosystem II, which

means that, upon illumination, the light-driven oxidation of PSI is

compensated for by its PSII-mediated reduction by electrons extracted from

water. This is shown in the fig. 1, where P700 reaches a redox steady state

where the PSI centers are now a mixed population having their primary donor

either in its reduced or oxidized states, the proportion of which depends on

the light intensity used. We also needed to block the electron flow from PSII in

order to quantify the original redox state of the PQ pool – however, in this

case we are in possession of a potent inhibitor 3-(3,4-dichlorophenyl)-1,1-

dimethylurea, DCMU, which blocks electron transfer out of PSII right after its

primary quinonic acceptor QA.

122

Fig. 1. Typical traces of P700 oxidation in Chlamydomonas in the presence or

absence of DCMU (100 µM here and throughout). The curve is a result of

subtraction of absorption kinetics at 730 nm (PC contribution) from 705 nm

changes (PSI and PC redox changes contribution). Light intensity – 100 e-

/s/PSI. Inset – a zoom on the reduction of P700+ in darkness following the

illumination.

Being a quinone analogue, DCMU binds with high affinity to the quinone-

binding pocket of the PSII, thus replacing the QB quinone and preventing the

transfer of electrons from QA to its acceptor. Acting rapidly and in sub-

millimolar concentration range it is more than suitable for the purpose stated

in the paragraph before. As shown in the figure 1, when 100 M DCMU is

used, a stable, much more complete oxidation of PSI is achieved.

However, does this minimal level correspond to 100% of the PSI centers with

an oxidized P700? DCMU blocks photogenerated linear electron flow from

water to PSI and thus decreases its reduction state. In case of a reducing

activity upstream of PSI, but independent of PSII, the addition of DCMU would

not block this source of reducing power if it does not go through the quinonic

PSII acceptor QA. This source of reducing power should result in an

underestimation of the total quantity of P700+ because a fraction of the

centers would stay reduced in the steady-state.

In steady-state conditions there are no changes of the redox state of the

P700:

123

𝑘𝑂𝑋[𝑃700𝑅𝐸𝐷] = 𝑘𝑅𝐸𝐷[𝑃700

𝑂𝑋 ]

or

[𝑃700𝑅𝐸𝐷] =

𝑘𝑅𝐸𝐷[𝑃700𝑂𝑋 ]

𝑘𝑂𝑋

Thus the concentration of P700+ will increase with the increase of the rate of

oxidation. This is easily obtained by increasing the light intensity, which

determines the kOX. Using a saturating pulse (5000 E/m2/s) at the end of the

illumination sequence (300 E/m2/s) increases the quantity of oxidized P700

(Fig. 2). Although PSI would be fully oxidized only in infinite light, the error on

the maximal amount P+ remains negligible, because of the low activity of

PSII-independent reducing pathways in the presence of DCMU as previously

observed (REF)..

Fig. 2. Typical traces of P700 oxidation in Chlamydomonas in the presence of

DCMU. The curve is a result of subtraction of absorption kinetics at 730 nm

(PC contribution) from 705 nm changes (PSI and PS redox changes

contribution). Light intensity – 100 e-/s/PSI. Inset – a zoom on the effect of the

saturating pulse (first detection point after the pulse is the lowest absorption

detected) and of the reduction of P700+ in darkness.

Importantly, because linear electron flow – and so the majority of the reducing

flux towards PSI – is blocked by DCMU, and P700 is mostly oxidized, the use

of a saturating pulse does not result in a transient over-reduction of the

124

acceptor side of PSI. This is crucial because in the case where PSI electron

acceptors are largely reduced (Fd, NADPH), oxidation of the special pair

results in a rapid back-reaction from the primary acceptors of PSI

(phylloquinones, FX, FA and FB iron-sulfur clusters) (Amunts et al, 2010;

Brettel, 1997; Rutherford et al, 2012). Because this reduction pathway is

rapid, part of the centers appear to be in a reduced state when measured at

705 nm, as is the case in anoxia as shown in blue in the fig. 3 (Clowez et al,

2015; Takahashi et al, 2013). This process has been described earlier as a

“PSI acceptor side limitation” that may lead to an underestimation the actual

number of PSI centers present in a sample (Klughammer & Schreiber, 1994).

Fig. 3. A typical trace of P700 oxidation in Chlamydomonas in the presence of

DCMU in oxic or anoxic conditions. The graph is a result of substraction of

absorption kinetics at 730 nm (PC contribution) from 705 nm changes (PSI

and PC redox changes contribution). Light intensity – 70 e-/s/PSI. Red box –

illumination, black box – darkness. Red arrow – saturating pulse. Inset – a

zoom on the reduction of P700+ in darkness following the illumination.

Adapted from (Takahashi et al, 2013).

What is the state of the electron transfer chain before an illumination?

Because of their high mid-point potential, the special pair P700, plastocyanin,

cyt. f and the Rieske protein are reduced in darkness. The situation is less

clear-cut for the plastoquinone pool. In the case of higher plants it is virtually

completely oxidized in the dark, but in Chlamydomonas it has been reported

125

that up to 30% of the pool remains in reduced state in fully oxic conditions in

darkness (e.g. Houille-Vernes et al, 2011). The size of the PQ pool is of about

10 to 20 electron equivalents (5-10 quinones) per photosystem II.

What is the sequence of events starting at the onset of illumination in the

presence of DCMU? PSI becomes photo-oxidized, but it is rapidly reduced by

its the secondary donors that have more reducing midpoint potentials. P700

thus stays reduced until all electrons from the donor side are transferred to

the acceptor side of PSI. Then, it becomes oxidized. However, due to some

heterogeneity in the number of electron donors to which each PSI has access,

there is not sharp change in the oxidized signal at 705 nm but a progressive

evolution after a lag period.

A typical trace of the situation before attaining the steady state in the light is

shown in the top panel of the figure below:

Fig. 4. A typical trace of P700 oxidation in Chlamydomonas in the presence of

DCMU, with the transitory period (lag) zoomed in. The curve is a result of

126

subtraction of absorption kinetics at 730 nm (PC contribution) from 705 nm

changes (PSI and PC redox changes contribution). Light intensity – 100 e-

/s/PSI. Top, traces of illumination of a dark-adapted sample or a sample

oxidized 9 seconds earlier (also, see sections below). Bottom, the same but

the green area represents the surface integrated for the sake of our

measurements.

Our idea for the quantification of the redox state of the PQ pool was the

following: we measured the area below the oxidation curve of P700. The lag

in the oxidation should depend on the kOX (a product of light intensity and PSI

antenna size) and the quantity of electron donors available upstream PSI.

Therefore, assuming that we use the same light intensity and sample

concentration between samples, and adjust the antenna size in conditions - or

mutants- susceptible to exhibit differences, we could highlight differences in

the quantity of electrons in the PQ pool per one PSI unit.

In order to achieve that, as shown in the fig. 4 B, we calculate the area below

the P700 oxidation curve, between the moment the light was turned on and

the quasi-steady state of the oxidation, between approximately 300 and 800

milliseconds.

We chose to use a relatively strong light to be in a steady state close to fully

oxidized PSI. As shown in the fig. 5, in lower light intensities when the kOX is

not high, the non-PSII-dependent reducing pathways are important enough to

keep a substantial part of PSI reduced. This should be avoided because a

significant part of the PSI signal would be lost; also, the lower the light

intensity the longer it takes to attain a steady-state, during which time

unwanted regulation phenomena can occur. Moreover, as will be shown later,

strong light allows saturation of the electron flows and therefore estimation of

the maximal rates of the enzyme-driven reducing pathways. The use of high

light intensities comes without the drawback of losing temporal resolution,

because the lag in P700 oxidation is long enough to be correctly resolved with

multiple detection points along the curve. Furthermore, because the actual

quantification of the redox state of the PQ pool takes place in the sub-second

time range (typically 300-500 ms), the chlororespiratory flux is negligible in

127

this period yielding results virtually independent of the activities of PTOX and

NDA2 during the experiment. Finally, the signal-to-noise ratio of the JTS (for

P700, I/I of 3-6.10-3 between the reduced and oxidized forms, noise of

approx. 5.10-5) is so high that a single scan yields readily quantifiable curves.

Fig. 5. Typical traces of P700 oxidation in Chlamydomonas in the presence of

DCMU. The curves are a result of subtraction of absorption kinetics at 730 nm

(PC contribution) from 705 nm changes (PSI and PC redox changes

contribution). Light intensity – 2 (black curve) to 250 e-/s/PSI (violet curve).

Adapted from (Alric et al, 2010b). Top, whole scan. Bottom, zoom on the dark-

to-light or light-to-dark transitions.

Using the WT and the DPTOX2 to compare the areas below their P700

oxidation curves only confirms that the lack of PTOX results in an

overreduction of the electron transfer chain. To get quantitative values, we

used a mutant lacking plastocyanin, the donor of electrons for P700+.

Because the oxidation of P700 in this strain generates a stable, long-lived

128

radical of the special pair which is only very slowly reduced as it lacks its

secondary electron donor, one can compare its area below the PSI oxidation

curve with the areas obtained using the WT or other mutant strains in order to

express them as a number of PSI turnovers upon steady-state oxidation.

However, the information provided by a sole comparison between WT and

PTOX2 strains still remains limited – even if we obtain a precise number of

electrons present in the chain for both of these strains. If we can indeed

reveal differences in the redox steady-state of the PQ pool in different

physiological conditions, no information about chlororespiratory fluxes will be

available. This can be exemplified in a situation where the chlororespiratory

activity of both PTOX and NDA2 is upregulated 10 times in a given condition

and the NDA2 activity is not limited by the presence of its substrate. In this

case, the redox steady state of the pool would not change – in the steady

state:

𝑘𝑂𝑋[𝑃𝑄𝐻2] = 𝑘𝑅𝐸𝐷[𝑃𝑄], so

𝑘𝑂𝑋

𝑘𝑅𝐸𝐷=

[𝑃𝑄]

[𝑃𝑄𝐻2]

We have therefore developed a modified protocol to access at least one of the

kinetic constants.

In order to obtain the maximal rate of an enzyme, one needs to provide a

substantial quantity of substrate so that the enzyme is limited by its catalytic

rate and not by the abundance of the substrate. We reasoned that, by

oxidizing the PQ pool completely we could obtain the rate of the NDA2 by

exploring the redox state of PSI acceptors at different time points of dark-

adaptation after the illumination. The protocol followed a simple scheme and

is shown in the fig.: 6

1) In the presence of DCMU, the dark-adapted sample was illuminated by

strong light (marked by yellow boxes). This treatment oxidized P700

and all PSI secondary donors.

129

2) The light was turned off in order to allow chlororespiratory enzymes to

poise the redox state of the PQ pool for up to a minute. Because the

pool was oxidized by the illumination, it was reduced by

chlororespiration (NDA2) in darkness.

3) The light was once again turned on after different times of the dark

period for the purpose of quantification of the state of PSI donors using

the PSI oxidation lag, as marked by the red rectangle. The longest

period between the two illuminations was the one that did not yield any

more changes in the area for the WT, i.e. the result was comparable

with a dark-adapted sample. The shortest period we used, 1 s,

corresponds to a time necessary to largely reduce P700+.

Fig. 6. A typical trace of the double P700 oxidation in Chlamydomonas in the

presence of DCMU. Light intensity – 100 e-/s/PSI. Yellow boxes – illumination,

black boxes – darkness.

130

Results

Below is the result of our comparison of the reservoir of reducing species for

P700+ as a function of time in darkness between the PTOX mutant and a

control strain. The area under the P700 oxidation curve is plotted as a

function of the time between two illuminations. The latter was explored in the

1 second to 1 minute timescale, as for the WT there were no more changes if

the time was increased.

Fig. 7. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU. A first illumination was applied to oxidize the chain,

and a variable time in darkness, plotted on the x-axis, followed. The area

under the lag of PSI oxidation upon second illumination was then divided by a

corresponding area of a PC-lacking mutant, yielding a result in PSI turnovers.

Light intensity – 450 e-/s/PSI. The value for the dark-adapted sample is shown

at the “infinite” time of the scale.

Shortly after the first illumination, the area is small in the WT strain. This is

consistent with the fact that at first we oxidize the whole chain, and by not

leaving enough time for it to re-reduce, the oxidation of P700 is rapid. Initially,

during a first phase, the longer is the time in darkness, the bigger the area

becomes, which corresponds to the reducing activity. However, after the peak

131

is reached at about 9 seconds, the area becomes smaller until it reaches a

steady-state after around 40 seconds.

In the PTOX2 mutant, however, the shape of the plot is very different. The

peak of the area size is higher, and it decreases much more slowly than in the

WT. It was then tempting to attribute the first phase, corresponding to an

increase in the number of PSI turnovers (i.e. dt of 1 to 9 s) to a reduction of

the chain by NDA2, and the second phase to the oxidative activity of PTOX.

At this stage it is unclear why the curves exhibit a biphasic shape instead of

having a monotonous, logarithmic shape, which would be expected if both

those enzymatic activities were limited by the rate of the enzymes.

These features are highly reproducible and appear even in lower light

intensities than those used before, as shown below in the fig. 8.

Fig. 8. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU. A first illumination was applied to oxidize the chain,

and a variable time in darkness, plotted on the x-axis, followed. The area

under the lag of PSI oxidation upon second illumination was then divided by a

corresponding area of a PC-lacking mutant, yielding a result in PSI turnovers.

Light intensity – 100 e-/s/PSI.

132

However, the most striking observation, apart from the shape of the curve, in

this experiment is the very large number of PSI turnovers reached by each

strain. As outlined before, it is known that there is not more than 20 electron

equivalents downstream PSII that may act as donors to PSI. In addition, only

in anoxia, where PTOX is inhibited, is this value attained due to the full

reduction of the PQ pool. Therefore, how is it possible that the above

experiment delivers up to 100 PSI turnovers in the fully oxic conditions we

used? How can this phenomenon be, moreover, light intensity-dependent?

We thought that two processes can increase the apparent number of PSI

turnovers: highly active cyclic electron flow around PSI and charge

recombination occurring because of a temporary overreduction of the PSI

acceptor side.

Fortunately, it is fairly easy to test whether CEF and the back-reactions

contribute to the increased turnover number. We opted to use methyl viologen

(paraquat) as an exogenous electron acceptor of PSI. The rationale is that

primary PSI acceptors with relatively low potentials, such as FA or FB, as well

as soluble ferredoxin, would be oxidized by methyl viologen before being able

to transfer electrons back to the PQ pool, and at the same time prevent the

temporal overreduction. However, MV would not influence the steady-state of

chlororespiration because its potential is lower than that of the

NADP+/NADPH couple, therefore it would not oxidize NADPH. We anticipated

that, by using MV, we would suppress the CEF-reinjection of reductants at the

donor side of PSI, and charge recombination due to the acceptor side

limitation - both resulting in an increase of the oxidation lag. We thought this

will occur without an important alteration of the redox state of the chloroplast,

as the addition of MV would be performed in darkness before the experiment.

A titration of the P700 oxidation curves (in low light intensity) with methyl

viologen is presented in the fig. 9

133

Fig. 9. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU and varying concentrations of MV. A first illumination

was applied to oxidize the chain, and a variable time in darkness, plotted on

the x-axis, followed. The area under the lag of PSI oxidation upon second

illumination was then divided by a corresponding area of a PC-lacking mutant,

yielding a result in PSI turnovers. Light intensity – 100 e-/s/PSI.

The amplitude of the curve dramatically decreases with the increase in MV

concentration, together with a loss of the “overshoot” form of the graph

(compare with the previous figure). Concentrations of the drug above the 100

µM range are necessary in order to decrease extensively the number of PSI

turnovers. The curve corresponding to the highest MV concentration

resembles strongly an asymptotic function, which is expected for the

aforementioned two-enzyme, rate-limited reaction, as shown below. We

continued to use 6 mM MV in order to be in a saturating condition.

134

Fig. 10. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU and 6 mM MV. A first illumination was applied to

oxidize the chain, and a variable time in darkness, plotted on the x-axis,

followed. The area under the lag of PSI oxidation upon second illumination

was then divided by a corresponding area of a PC-lacking mutant, yielding a

result in PSI turnovers. Light intensity – 450 e-/s/PSI.

Fig. 11. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU or DCMU and MV (6mM). A first illumination was

applied to oxidize the chain, and a variable time in darkness, plotted on the x-

axis, followed. The area under the lag of PSI oxidation upon second

135

illumination was then divided by a corresponding area of a PC-lacking mutant,

yielding a result in PSI turnovers. Light intensity – 450 e-/s/PSI.

We can thus conclude that the use of MV yields essentially the result we

expected at the beginning of the project – a measure of the NDA2-mediated

reduction of the PQ pool. The number of electrons present in the PSI donors

is on its low side in these measurements, yielding 4, however it varies

between 4 and 8 for the WT, which is roughly consistent with the expected

number of PSI electron donors.

In the PTOX2 strain the number of electrons available for PSI reduction is

higher than in the WT, as expected, and varies between 7 and 13. It confirms

the utility of this method in counting the PSI electron donors.

Fig. 12. Evolution of the area under P700 oxidation curves upon illumination in

the presence of DCMU and 6 mM MV in the PTOX2 strain. A first

illumination was applied to oxidize the chain, and a variable time in darkness,

plotted on the x-axis, followed. The area under the lag of PSI oxidation upon

second illumination was then divided by a corresponding area of a PC-lacking

mutant, yielding a result in PSI turnovers. Light intensity – 450 e-/s/PSI. Red,

exponential fit of the data. Inset, a zoom of the initial slope.

136

In the figure 12, a magnification of the reduction phase observed in the PTOX-

lacking strain is presented. We calculated the value of the maximal rate of

reduction of the electron transfer chain by estimating the initial slope of the

reducing activity. The slope of approx. 2 proves a reduction rate of 2 e-/PSI/s,

in perfect agreement with the maximal rate of NDA2 calculated using

fluorescence measurements (Houille-Vernes et al, 2011). This further

validates our method as a reliable way of measuring the redox activities in

photosynthetic chain.

However, the effect of methyl viologen is two-way. As outlined before, it is

responsible for decreasing the number of reduced CEF donors (i.e.

Ferredoxin), but also for accepting electrons from PSI. Therefore, in case of a

temporal over-reduction of the PSI acceptor side upon illumination, a fraction

of the P700 oxidation events would result in a rapid charge recombination

(see fig. 13) in the absence of MV but not in its presence, yielding a special

pair that is reduced by back-reactions and not through linear or cyclic

electron. To find out whether such contribution from back-reactions was

significant in our experimental procedure we used the classical saturating

pulse method (Klughammer and Schreiber, 1994) to evaluate the PSI

acceptor side limitation. Briefly, saturating pulses were superposed on the

actinic light during the lag period to investigate whether the increase in kOX

would yield a corresponding increase in P700+ concentration (CEF) or not

(presence of the acceptor side limitation).

137

Fig. 13. Structure of PSI electron carriers and the halt-times of charge

recombination from a given reduced acceptor to P700+. Data from (Amunts et

al, 2010; Brettel, 1997; Rutherford et al, 2012).

Fig. 14 shows the traces of PSI oxidation during the lag period, where the

saturating pulses (5000 E/m2/s, roughly 3000 e-/s/PSI) were superimposed

on the actinic light. It is visible that no matter the precise time the pulse was

applied (at 80 or 150 ms of illumination), or its duration (2-50 ms), PSI was

only partly photooxidizable by pulses during the lag (the total extent being

similar to the steady-state value in the right-side of the figure). This suggests

that a part of the surplus area we are measuring is due to back-reaction from

primary PSI acceptors and part from cyclic and linear electron flows (because

if solely the CEF was the reason, its rate would exhibit roughly 4000 e-/s/PSI if

40% of P700 was oxidized during a pulse (see figure below), value which is

inconceivable).

138

Fig. 14. A typical trace of P700 oxidation in Chlamydomonas in the presence

of DCMU, with the transitory period (lag) zoomed in. Light intensity – 100 e-

/s/PSI throughout, but with saturating pulses (4000 e-/s/PSI) of different

lengths (6 or 40 ms) were superposed at 50 or 135 ms of illumination

(arrows).

This observation is consistent with the effect of methyl viologen, as use of this

drug would simultaneously competitively inhibit CEF and alleviate any PSI

acceptor side overreduction by providing large amounts of electron acceptor.

We have hence established that by using methyl viologen we monitor,

by the mean of the lag phase, the electron flow from PQ pool to PSI, and

without the drug we explore a convolution of three activities. First,

inefficient PSI turnovers, in centres with overreduced acceptor side,

which lead to charge recombination and not to an oxidation of donors

initially present in the membrane. Secondly, two types of actual PSI

turnovers – efficient, when PSI oxidizes the pool of donors and transfers

them through the ‘linear’ route; and cyclic-type PSI turnovers, in the

case of electrons that come back to the membrane after having been

transferred to the stroma.

139

Obviously, in the next step wanted to discriminate between the CEF and

back-reaction contributions to the surplus area of PSI oxidation. It is

interesting to understand why this phenotype has its peak within the first 9

seconds of darkness between the two illuminations, and whether it is due to i)

transfer of a few electrons from the PQ to the NADPH pool upon the first

illumination, which would then result in PSI over-reduction during the next

illumination, or ii) changes in CEF activity triggered by the first illumination. In

order to discriminate between the two hypotheses, we developed a

completely novel protocol.

Distinguishing back-reactions from cyclic electron flow

Measurements of the redox state of P700 do not discriminate between the

reduction pathways, i.e. both the PC-mediated reduction and back-reactions

will result in identical changes in the absorption of the special pair. One could

imagine discriminating the two processes with an appropriate time-resolution.

Yet this is a difficult task because, upon light-to-dark transition, a pool of PC+

and P700+ will be simultaneously reduced both by the b6f and by back-

reactions, the rate of the former process depending on the size of the PC

pool, on how much PC is pre-reduced and pre-bound to PSI, and on the

quantity of P700 reduced in the light. As shown in the figure 13., the reduction

by charge recombination from reduced A0, A1 and FX to P700+ is virtually

achieved within 1 ms, whereas back-reaction from FA/FB is much longer, with

a half-time of 50 ms. These timescales partly overlap with the PC-mediated

reduction of P700, yielding the analysis extremely complex.

We therefore proceeded to measure the photochemical rate using the DIRK

(Dark Interval Relaxation Kinetics; (Joliot & Joliot, 2002; Sacksteder &

Kramer, 2000)) method in order to exclude the back-reactions from the

evaluation of photochemical rates.

As previously described (Joliot & Joliot, 2002; Sacksteder & Kramer, 2000),

the DIRK method allows a quantification of the instantaneous photochemical

rate of the photosystems – or, in our case, of PSI because we inhibit PSII in

our experimental procedure. DIRK will be described here, yet it is first

necessary to understand the principles of the physical phenomenon, which is

140

used by DIRK. Electrochromic shift (ECS) experiments consist of detecting

changes in the electrochromic signal during the transition from light to dark.

The mechanism of this signal is the following: during illumination, an electric

field is formed across the thylakoid membrane due to the activity of

photosystems and stroma-to-lumen proton displacement (thanks to the PSII-

PQ-cyt. b6f series of reactions). This electric field modifies the electronic

properties of some pigments embedded in the membrane, shifting their

absorption spectrum in what is called a Stark effect, yielding an easily

measurable and quantifiable signal.

Fig. 15. Electrochromic shift. A, a schematic representation of the ground and

excited state of a chromophore in the presence or in the absence of an

electric field. B, Changes in the absorption spectrum of a pigment induced by

an external electric field, and a difference spectrum (field-no field). Adapted

from (Bailleul et al, 2010).

We will now proceed to introduce the DIRK method to the reader. During

steady-state illumination, the electric field is produced by the aforementioned

141

enzymes, and consumed by the ATP synthase and other “leaks”, and its rate

of change is 0 (corresponding to a flat ECS signal). In the light:

𝑅𝐿 = 𝑅𝑃ℎ + 𝑅𝑏𝑓 − 𝑅𝑙𝑒𝑎𝑘, where the 𝑅𝐿 is the rate at which the electric

component changes in the light, 𝑅𝑃ℎ is the photochemical rate, 𝑅𝑏𝑓 the

turnover of the cyt. b6f, and 𝑅𝑙𝑒𝑎𝑘 the rate of the field decay.

Fig. 16. A schematic representation of the ECS signal changes and the

theoretical considerations of contributions to the signal upon illumination and

cessation of illumination. Adapted from (Bailleul et al, 2010)

As outlined by Joliot (Joliot et al, 2004a; Joliot & Joliot, 2002), upon cessation

of the illumination, the photochemical rate falls to zero in sub-microsecond

timescale after excitation relaxation and charge separation, but the b6f

continues its light-independent turnover in the millisecond time range. The

leak is also initially unchanged because although the rate of ATP synthase

depends on the electrochemical gradient, its change in this timescales is

negligible. Therefore, in darkness after the light-to-dark transition:

𝑅𝐷 = 𝑅𝑏𝑓 − 𝑅𝑙𝑒𝑎𝑘, where 𝑅𝐷 is the experimentally measured rate of decay.

Therefore,

𝑅𝐿 − 𝑅𝐷 = 𝑅𝑃ℎ + 𝑅𝑏𝑓 − 𝑅𝑙𝑒𝑎𝑘 − (𝑅𝑏𝑓 − 𝑅𝑙𝑒𝑎𝑘) = 𝑅𝑃ℎ,

or in the steady-state illumination where 𝑅𝐿 = 0: −𝑅𝐷 = 𝑅𝑃ℎ

142

The slope, in order to provide absolute values, must be however normalized.

This is done using a saturating, single turnover laser flash in the presence of

HA and DCMU, which block PSII turnover. The flash intensity is high enough

to drive a charge separation in virtually all photosystem I in the sample, but its

duration is short enough to allow only a single photochemical even in each

PSI center, hence it provides the electrochromic shift value for “one positive

charge in the lumen per photosystem” (with the inhibitors, one positive charge

per PSI). In terms of the ECS signal, the photochemistry creates a positive

charge on the lumenal side and a negative charge in the stromal side of the

membrane – in turn provoking the Stark effect in some of the membrane

pigments whose orientation is always fixed with regard to the membrane. This

polarization is detectable typically 100 s after the flash as this time is long

enough for excitation transfer and charge separation, but short enough to

capture a situation before the cyt. b6f starts its turnover or the ATP synthase

starts to use the electric field, dissipating it. We can hence divide the slope by

the value of the ECS of the same sample upon a saturating flash in the

presence of PSII inhibitors:

𝑆𝑙𝑜𝑝𝑒@1−5𝑚𝑠

[∆ (∆𝐼

𝐼⁄ )

𝑠]

𝐹𝑙𝑎𝑠ℎ𝐷𝐶𝑀𝑈,𝐻𝐴 [(∆𝐼

𝐼⁄ ) 𝑒−

𝑃𝑆𝐼⁄]

= 𝑅𝑒𝑑𝑢𝑐𝑡𝑖𝑜𝑛 𝑟𝑎𝑡𝑒1−5𝑚𝑠[∆𝑒−

𝑠 ∙ 𝑃𝑆𝐼 ]

Obtaining the photochemical rate of PSI at any given time.

A unique advantage of employing the ECS-based measurements is that

the signal remains if P700+ is reduced by PC, yet it will detect a charge

recombination-type of reduction since the electric field that produces

ECS collapses upon charge recombination. The latter occurs since the

transfer of an electron in the membrane from the stromal position to the

special pair located in the lumenal part of it is electrogenic, i.e. yields a

(negative) electrochromic signal due to the movement of the negative charge

to the positive (p) side of the thylakoid membrane. Meanwhile, PC-mediated

reduction does not result in a change of the signal in short timescales,

143

because a positive charge in the lumen side of the membrane is simply

moved to the PC, also located in the lumen, upon its oxidation by the P+.

The back-reaction is virtually finished at 1 millisecond as shown in the figure

from Joliot & Joliot (2004):

Fig. 17. Changes in membrane electric potential in darkness after an

illumination. 1, conditions of PSI acceptor side limitation. 2, conditions where

the PSI acceptor side limitation is alleviated. Adapted from (Joliot et al,

2004a).

Hence, by measuring the ECS decay slope between 1 and ~5 ms after the

light is turned off, we can obtain information about the true photochemical rate

of PSI (while blocking PSII acceptor side with DCMU and providing

hydroxylamine, which is a potent reductant of P680+) without contribution from

the back reactions.

How does measuring an instantaneous photochemical rate of photosystems,

however, helps to distinguish between CEF, LEF and back reactions

throughout our at-least-500 ms protocol determining the redox state of the

P700? It is necessary to multiply the DIRK measurements in order for them to

span the whole time course of PSI oxidation, until a redox steady-state is

attained. We thus designed a multiple DIRK experiment to follow the course of

oxidation in the following manner:

144

Fig. 18. Typical traces of P700 oxidation in Chlamydomonas in the presence

of DCMU, with the transitory period (lag) zoomed in. The curve is a result of

substraction of absorption kinetics at 730 nm (PC contribution) from 705 nm

changes (PSI and PS redox changes contribution). Light intensity – 100 e-

/s/PSI. The two traces of oxidation represent a dark-adapted sample or a

sample oxidized 9 seconds earlier Grey vertical lines represent moments (i)

where the light was turned off in order to monitor the rate of decay of the

membrane potential with the DIRK method. i values: 10, 20, 30, 50, 75, 150,

300, 450, 750, 1500, 2000, 3000, 4000, 5000 ms.

We placed the light-to-dark transitions allowing the assessment of the

photochemical rate along the kinetics of PSI oxidation at the time points

marked by vertical lines in the figure 18, and beyond them, until the end of the

light period. Each dark period lasts for 6 ms in order not to perturb the system

with excessive dark periods. Because of the density of the measuring points

at the beginning of the oxidation period, the protocol was divided in two steps,

with a first measurement at points i, i+2, i+4 etc. followed by a second

measurement at points i+1, i+3… . With this method, the dark interval length

was negligible with regards to the period in the light and the illumination could

be considered as continuous.

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Similarily, in order to elucidate the dependence of the changes in

CEF/LEF/charge recombination pathways leading to a reduction of P700

during the dark period between two illuminations, this experiment was

conducted not only for the dark-adapted sample but also spanned through the

range of dt used earlier for the 705 nm absorption changes measurements.

The time points chosen to best resolve the reduction processes were: 1, 5, 9,

15 and 35 s of darkness, and the DIRK measurements were employed both

upon the first and second illumination. The first illumination represents a dark-

adapted state and will be further displayed as the dt=60 s in order to facilitate

the graphical representation.

For a given light, upon a dark-to-light transition in the presence of PSII

inhibitors, the initial rate (when all PSI centers are open) of the ECS signal

rise (measured when no more than 20% of the centers undergo charge

separation, i.e. up to 1-2 ms) will be proportional to the PSI antenna size at

the time of illumination, allowing correction for possible differences in antenna

between strains. In the present case, this is of minor importance because,

even though the PTOX2 mutant is in state II when dark-adapted, i.e. its PSI

antenna size increases with regards to that in the WT, the light intensity we

used, as above for the P700 measurements, is close to saturation compared

to the maximal rate of the non-photochemical processes (i.e. b6f turnover).

However, the initial maximal rate of photosystems can be used as well to

precisely determine the nominal light intensity in a similar way that we used

during the light-to-dark transition – by dividing the slope by the saturating,

single turnover flash value. Since upon the dark-to-light transitions the light

intensity determines the maximal possible photochemical rate because the

centers are all open, it will be reported in the following graphs at the time

t=1ms of the illumination.

The results are presented below:

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Fig. 19. Results of the multiple DIRK experiment in the WT. Each value of the

electron flow through the PSI during a 5 second illumination is represented as

a column. i values (moments when the PSI ETR was measured): 10, 20, 30,

50, 75, 150, 300, 450, 750, 1500, 2000, 3000, 4000, 5000 ms. dt (time in

darkness after a first oxidation) values: 1, 5, 9, 15, 35 s and dark adapted

(shown at 60 s). Each value is an average of 8 technical replicates each for 2

biological replicates, apart from the dark-adapted value, which is an average

of 40 technical replicates each of 2 biological samples.

Roughly, it is qualitatively visible that the initial rate of PSI quickly (within 1 s)

decays to a small value, consistently with the previous PSI redox

measurements. It can be also seen that the intermediate dt yields in general

higher values than the dark-adapted sample, and that for the dt=1 the rates

are by far the slowest (also in agreement with the very small values for P700

lag area values we reported before). The latter is due to the low quantity of

electrons in the chain after 1 second of darkness.

Because the graph with the entire data acquired during the experiment is not

completely readable, we started by calculating the integral of the curve

connecting each measurement point until the steady-state, in a similar way

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that was used before for the 705 nm measurements. Because the value of

instantaneous PSI photochemical rate is expressed in e-/s/PSI, its integration

over time yields number of electrons transferred by PSI until it reaches the

nearly-oxidized redox steady-state.

Fig. 20. Result of the multiple DIRK experiment for a dark-adapted WT strain.

Gray area represents the surface, which was integrated; its value is presented

above (“Area”) and expressed in milielectrons. Red line – baseline of the

integration, which value is an average of the last 5 measuring points (i.e.

steady-state). Each point is an average of 2 biological replicates, each

measured 40 times.

The integrated area for the dark-adapted wild-type strain is shown in the

figure above in grey. The number of electrons transferred in this case is

slightly more than 15 per PSI. This is both significantly less than what was

inferred from P700 redox measurements in the same light intensity (over 30),

and more than the number of electrons in PSI donors in darkness (between 4

and 8). Because we eliminated the former overestimation thanks to the

employment of the DIRK method instead of P700, and the number of

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electrons transferred by photosystem I is higher than the number of available

electrons from the linear flow, it is necessary to admit that a rapid reducing

pathway between PSI and PSII is involved in injecting electrons in the

intersystem electron carriers. Clearly this process corresponds to electron

flows much larger than those attributed to chlororespiration (Houille-Vernes et

al, 2011), therefore preventing us to study properly the latter pathway with this

experimental procedure.

An obvious candidate to account for the surplus electrons is cyclic electron

flow. This elusive process is responsible for the reinjection of electrons from

reduced Ferredoxin back to plastoquinones. It is usually considered that there

are two types of CEF: the faster, pgr5/pgrl1-dependent, and the slower NDA2

(or NDH in the case of plants)-dependent. The latter would use NAD(P)H as a

substrate in algae, whereas the pgr5/pgrl1-dependent is rather thought about

as a ferredoxin:plastoquinone oxidoreducing activity. However, it was already

shown that the absence of pgr5 does not lower the maximal rate of CEF in

plants (Nandha et al, 2007) proving a regulatory role and not a direct

component of CEF. This is apparently also the case for pgrl1 (P. Joliot,

personal communication). Hence, throughout the text I will refer to the

NDA2 as having a chlororespiratory activity, and to the rapid donation of

electrons from the stroma to the membrane simply as CEF, whether or

not it is regulated by pgr5/pgrl1.

In our measurements, upon oxidation of PSI and transfer of electrons to the

stroma, ferredoxin would become reduced and able not only to reduce NADP+

thanks to the FNR, but also to return some electrons back to the chain. This

activity would need to be only transient, as after the initial lag, P700 is not

being reduced in a rapid way (e.g. see the “Initial idea overview –

measurements of chlororespiratory fluxes“ section).

The integrals from each dt can be presented on the same graph as the P700

data:

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Fig. 21. Comparison between the evolution of the number of PSI turnovers

before complete oxidation as measured with DIRK or area-under-P700-

oxidation-curve methods in the presence of DCMU. A first illumination was

applied to oxidize the chain, and a variable time in darkness, plotted on the x-

axis, followed. The area below the curve from connected points expressing

instantaneous PSI ETR has been integrated and is shown here for the WT

(blue) and the PTOX mutant (orange) for each dt. The area under the lag of

PSI oxidation upon second illumination was divided by a corresponding area

of a PC-lacking mutant, yielding a result in PSI turnovers for the WT (black)

and the PTOX2 (red). Light intensity – 450 e-/s/PSI in both experiments.

Multiple features of the integrated DIRK curves can be pointed out here in

comparison with the P700 experiments. First of all, the amplitudes are, as

discussed before, much lower for both the mutant and the WT. The form of

the curves is also altered, being flattened in both cases, yet still conserving

the maxima around 9-15 s of darkness before re-illumination. Using DIRK, the

PTOX differs from the WT mostly in amplitude, and not in shape, contrary to

the P700 method. Yet, especially for the long dark-adaptation times, one can

see that the quantity of charge recombination is much higher in the PTOX

mutant with regards to WT by the simple mean of comparing the values that

include LEF, CEF and back-reactions (P700) or only CEF and LEF (DIRK).

150

This last observation is consistent with the observations we made earlier in

vivo, that both the PQ and the NAD(P)H pools are more reduced in PTOX2

which induces more PSI acceptor side limitation and thereof more back-

reactions contributing to P700 re-reduction.

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Implications for CEF mechanism

I wish here to further analyze the contribution of CEF upon illumination in

presence of DCMU. Although far more speculative because they depend on

several assumptions that seem logical but may not be true, the suggestions

below lead to a different type of thinking about cyclic when compared with

most of the recent literature about CEF in Chlamydomonas.

Let us return to the result of the DIRK experiment described before for the

dark-adapted wild-type:

Fig. 22. Result of the multiple DIRK experiment for a dark-adapted WT strain.

Gray area represents the surface, which was integrated; its value is presented

above (“Area”) and expressed in milielectrons. Red line – baseline of the

integration, which value is an average of the last 5 measuring points (i.e.

steady-state). Each point is an average of 2 biological replicates, each

measured 40 times.

The first approximation

In the saturating light we used, let us assume for the sake of simplicity that the

rate of electron flow, limited by the b6f, is of about 200 e-/s/PSI (1 electron

transferred every 5ms). As shown before, the algae originally have a partially

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reduced electron transfer chain and about 7 electron equivalents upstream

PSI (a number which may be overestimated). With this rate of electron

transfer, PSI should get oxidized within 35 ms if only linear electron flow was

operating. We see, however, that after 35 ms the rate of PSI still is very high

(~ approx. 120 e-/s/PSI). This prompted us to hypothesize that a fast, higher

plant-like CEF operates in Chlamydomonas regardless of the redox state of

the stroma, but the ability to detect it depends on the method used.

Development of the model

What are the events that unfold upon illumination in presence of DCMU? We

propose the following scenario:

1. In darkness, chlororespiration is in steady-state, constantly reducing

and oxidizing quinones. It was shown that in steady-state, the NDA2

reduces about 1 quinone/s/PSI (Houille-Vernes et al, 2011), yet the

maximal rate of this enzyme in oxic conditions and fully oxidized PQ

pool (although perhaps limited by NAD(P)H quantity, still corresponding

to our experimental setup) is of 2 e-/s/PSI. Because the ferredoxin pool

is virtually completely oxidized, CEF is inactive. The pool of electron

donors consists of about 7 e-/PSI, as evidence by P700 experiments in

the presence of DCMU and MV.

2. At the onset of illumination, PSI works at very high rates, as

determined by the rate of PC-mediated P700+ reduction, or limited by

the light intensity.

Meanwhile, cyclic electron flow is not substrate-limited because the

acceptor side of PSI is over-reduced as evidenced by the P700

measurements. The presence of reduced ferredoxin means that CEF

can start to transfer electrons back to the PQ. From the first

miliseconds of illumination, CEF and LEF are both working

simultaneously. Some of the electrons are not to be recycled, however,

and continue the linear pathway, for instance reducing NADPH or

oxygen (for simplicity, the “NADPH reduction” will be assumed

thereafter); those that are redirected to the thylakoid membranes are

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thus counted as a part of the initial electron donor pool as they are

taken into account by the DIRK measurements.

3. When the PC pool and the high potential cofactors of cyt. b6f, which

were reduced in darkness, are fully oxidized, the rate of PSI electron

transfer is determined by the slowest step of the electron flow, which is

the oxidation of plastoquinones at the QO site of cyt. b6f. Because the

quinol oxidation becomes less and less energetically favourable with

the acidification of lumen (as it includes proton release to the lumen),

this rate is difficult to predict, but its value should not be lower than 150

e-/s/PSI.

4. Throughout the illumination, because of the absence of PSII activity,

the initial pool of electrons that are being transferred back from PSI

acceptor side is becoming smaller and smaller due to a part of the

electrons reducing NADPH. At some point, the size of this pool starts to

limit the rate of PSI electron transfer instead of the rate of PQH2

oxidation – i.e. PSI is limited by the absence of its substrate, reduced

PC.

5. After about 400 ms, the electrons initially present in the membrane are

all transferred to NADPH, and PSI as well as its electron donors are in

their redox steady-state, close to complete oxidation.

6. Why is still a little part of PSI centres reduced? In presence of DCMU,

the only entry site for electrons to the membrane is the NDA2, its

activity sustains a non-zero ETR of PSI. However, the rate of PSI ETR

is larger than the maximal rate of NDA2 – we postulate, that similarily

to the previous situation, a part of electrons that were just transferred to

the Ferredoxin can return to the membrane via the cyclic electron flow,

therefore increasing the rate of PSI ETR (10 e-/s/PSI) with regards to

the maximal rate of NDA2 (2 e-/s/PSI). This proposition is essential for

the comprehension of this model – the NDA2 supplies CEF with

electrons in the presence of DCMU.

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Fig. 23. State of the electron carriers in the dark (A) and in the light in

presence of DCMU (B). The pools of carriers are presented as amphoras as

proposed in (Lavergne, 2009), with the area of P700 being equal to 1

electron. The thickest part of the amphora corresponds to the midpoint

potential of the carrier and is shown on the y-axis. Adapted from (Alric, 2010).

For a visual presentation, consider the scheme above, adapted from (Alric,

2010), which describes the steady-state redox status of the electron

transporters in oxic conditions in darkness and in the light in the presence of

DCMU. We postulate that in the transitory period, upon illumination, the

Fd/FNR pool becomes reduced (i.e. the red area is exchanged for the blue

area of Fd) and it can transfer electrons directly to the PQ pool with rates

comparable to those in higher plants (before the Fd transfers its electrons to

the NADPH pool). The pool then becomes oxidized but is re-reduced through

CEF, until too many electrons are transferred to the NADPH pool and are no

longer available for CEF.

Let us come back to the point 6. of the previous paragraph. We propose that

in the steady-state illumination in the presence of DCMU three processes are

operating simultaneously: slow reduction of the PQ pool by NDA2; CEF which

uses the electrons supplied by the NDA2 and which creates a ‘short-circuit’

from Fd back to the plastoquinones; and leak of electrons towards the linear

route, which is of course the reason for which not all of the electrons are

recycled. If this interpretation is correct, we can define the probabilities that an

155

electron just transferred to the stromal side of PSI is going to follow the CEF

or LEF route as shown in the figure below:

Fig. 24. A scheme of the electron transfer chain in the light and in the

presence of DCMU. Orange, thylakoid membrane. Blue, some of the pertinent

photosynthetic complexes, from the left PSII blocked with the DCMU, NDA2,

cyt. b6f and PSI. The rates of the NDA2-mediated PQ reduction (Houille-

Vernes et al, 2011) and of PSI reduction (e.g. Takahashi et al, 2013) are

shown. 𝑃𝐶𝐸𝐹, probability that an electron transferred by the PSI returns to the

membrane via a rapid cyclic process (i.e. FQR)

These probabilities are given by:

𝑃𝐶𝐸𝐹 = 1 − 𝑃𝐿𝐸𝐹, where 𝑃𝐶𝐸𝐹 is the chance that the electrons to re-reduce the

PQ and 𝑃𝐿𝐸𝐹 is probability that the electrons are not recycled.

𝑃𝐶𝐸𝐹 =𝑃𝑆𝐼 𝑟𝑎𝑡𝑒−𝑁𝐷𝐴2 𝑟𝑎𝑡𝑒

𝑃𝑆𝐼 𝑟𝑎𝑡𝑒, so in the case of fig. 24, 80%; i.e. in order to observe

a PSI rate of 10e/s, with a known electrons influx to the membrane of 2e/s,

each electron needs to be recycled on average 4 times before its is released

to continue along the linear pathway.

For the situation presented in the fig. 24, the measured rate of PSI ETR is 6 e-

/s/PSI, yielding a CEF probability of 66% assuming that upon oxidation of the

PQ pool, the NDA2 works at 2 e-/s/PSI as shown before (Houille-Vernes et al,

2011).

How to calculate the same probability during the transitory period of the PSI

oxidation, where the steady-state values are changing? Assuming that the

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probability of CEF stays identical throughout the 500 ms illumination, and that

every electron of the initial pool of electrons with the size x, which happens to

be transferred to the acceptor side of the PSI has a given 𝑃𝐶𝐸𝐹 and 𝑃𝐿𝐸𝐹, the

total quantity of electrons transferred byt the PSI is described by a geometric

series:

𝑁𝑢𝑚𝑏𝑒𝑟 𝑜𝑓 𝑒𝑙𝑒𝑐𝑡𝑟𝑜𝑛𝑠 𝑡𝑟𝑎𝑛𝑠𝑓𝑒𝑟𝑟𝑒𝑑 = 𝑥 + 𝑥 ∙ 𝑃𝐶𝐸𝐹1 + 𝑥 ∙ 𝑃𝐶𝐸𝐹

2 + ⋯ + 𝑥 ∙ 𝑃𝐶𝐸𝐹𝑛−1

Because the 𝑃𝐶𝐸𝐹 is smaller than 1, this series is converging and finite. In this

case the total number of electrons transferred, 𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾:

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 =𝑥

(1 − 𝑃𝐶𝐸𝐹)

As we know the actual initial number of electrons 𝑥 = 7, and the total number

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 = 15:

𝑃𝐶𝐸𝐹 =𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝑥

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾

𝑃𝐶𝐸𝐹 =15 − 7

15= 53%

53% is arguably smaller than 66%, although the rough range of the

probabilities stays the same. However, our initial hypothesis assumes that

𝑃𝐶𝐸𝐹 does not change throughout the illumination. Is this a possible source of

an error? The electrons reinjection to the membrane step is thought to be

limiting for this cyclic (although it has not been demonstrated) and let us

assume this is the case here.

For the sake of simplicity, let us propose that the maximal rate of CEF in

Chlamydomonas is the same as in plants, i.e. 120 e-/s/PSI, and that this

limiting rate is the rate of the Ferredoxin:plastoquinone oxidoreductase. What

does it change in our reasoning? At the beginning of the illumination, the rate

of PSI is higher than 120 e-/s, so only a part of the electrons transferred to the

PSI acceptor side are able to be recycled. The probability of them returning to

the PQ is therefore lower than in conditions where the rate is not saturating.

Because computation of dynamically changing probabilities in a geometric

157

series is difficult, we will inverse the rationale, and try to adjust the values of 𝑥

and 𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 so that they more precisely reflect what happens. If only a part of

electrons can be recycled, and the electrons lost due to a faster linear flow

than the CEF are 𝐿:

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝐿 =𝑥 − 𝐿

(1 − 𝑃𝐶𝐸𝐹)

or

𝑃𝐶𝐸𝐹 =(𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝐿) − (𝑥 − 𝐿)

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝐿

which reduces to:

𝑃𝐶𝐸𝐹 =𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝑥

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝐿

The electrons which are not available for cyclic are those given by the integral

of the DIRK measurements with a baseline of 120, our maximal rate of cyclic:

Fig. 25. Result of the multiple DIRK experiment for a dark-adapted WT strain.

Gray area represents the surface, which was integrated; its value is presented

above (“Area”) and expressed in milielectrons. Red line – baseline of the

integration, which value corresponds to 120 e-/s/PSI. Each point is an average

of 2 biological replicates, each measured 40 times.

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The integration with a baseline of 120 shows that 3,3 electrons per PSI should

potentially be excluded from the cyclic pathway; therefore:

𝑃𝐶𝐸𝐹 =𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝑥

𝐸𝑡𝑜𝑡,𝐷𝐼𝑅𝐾 − 𝐿=

15 − 7

15 − 3.3= 68%

The adjusted 𝑃𝐶𝐸𝐹 = 68% of CEF with a maximal rate of 120e/s/PSI is in

perfect agreement with the probability counted in the steady-state, when PSI

rate is limited by the quantity of reduced PC (66%).

Note that the data presented above, although perhaps not proving the

presence of the rapid cyclic, strongly points into this interpretation even for the

dark-adapted sample. The integral of the DIRK measurement for the dark

interval of, for example, 9s between illuminations suggests that the 𝑃𝐶𝐸𝐹 is

even higher (+50%) in these conditions and necessitates even more our

model of the CEF.

For the sake of the argument, one needs to refer these findings/hypotheses to

those reported earlier. It was shown that the maximal rate of CEF in oxic

conditions (as measured in presence of DCMU) was 10 e-/s/PSI. If indeed this

was the maximal rate, how would our calculations look?

Fig. 26. Result of the multiple DIRK experiment for a dark-adapted WT strain.

Gray area represents the surface, which was integrated; its value is presented

159

above (“Area”) and expressed in milielectrons. Red line – baseline of the

integration, which value is 10 e-/s/PSI. Each point is an average of 2 biological

replicates, each measured 40 times.

The adjusted 𝑃𝐶𝐸𝐹 value in this case (baseline of 10 e-/s/PSI) equals 470%, a

number that is impossible for a probability (always between or equal to 1 and

0).

If the maximal value of CEF was identical to the one found in anoxic

conditions (50 e-/s/PSI):

Fig. 27. Result of the multiple DIRK experiment for a dark-adapted WT strain.

Gray area represents the surface, which was integrated; its value is presented

above (“Area”) and expressed in milielectrons. Red line – baseline of the

integration, with a value of 50 e-/s/PSI. Each point is an average of 2

biological replicates, each measured 40 times.

𝑃𝐶𝐸𝐹 = 99,4% - or in other words, that each electron of an extremely small part

of initial PSI donor pool, (𝑥−𝐿)

𝑥= 0,7%, would need to be recycled on average

161 times to account for the apparent increase in the area if a maximal rate of

CEF of 50 e-/s/PSI was to be believed. This would mean that a very small part

of PSI centers is capable of operating in a practically only-CEF mode.

160

Although all of our measurements were performed in oxic conditions, one can

calculate the change in the 𝑃𝐶𝐸𝐹 from the anoxic data in the literature.

Assuming that the NDA2 activity measured in oxic conditions where the

NAD(P)H pool is only partially reduced is its maximal rate, depending on the

article (Alric, 2014; Takahashi et al, 2013; Tolleter et al, 2011) the CEF rate

scatterd from 40 to 60 e-/s/PSI, which translates to 𝑃𝐶𝐸𝐹 of 95 to 97%. If the

rate of NDA2 was underestimated in oxia because of substrate limitation and

the true VMax is higher, say 5 e-/s/PSI, the 𝑃𝐶𝐸𝐹 drops to 88 to 92%. In both

cases, the multiple-fold increase of the CEF rate in anoxia with regards to oxic

conditions is explained by a relatively modest increase in 𝑃𝐶𝐸𝐹, expected

when the reducing pressure increases and the chloroplast stroma becomes

reduced.

We have finally pursued another way of convincing ourselves that the use of

𝑃𝐶𝐸𝐹 from the steady-state can be justified during the transition periods. In

order to do that, we reasoned that a strong indication of such possibility would

be if we could correlate changes in 𝑃𝐶𝐸𝐹 with changes in CEF quantity

measured during the transitions – i.e. their correlation would point us in a right

direction with regards to the theory.

Fortunately, we found indications that there could exist instances where the

𝑃𝐶𝐸𝐹 changes occur. They were found at the beginning of my PhD - as it often

happens, by chance - and in fact shown even before (Alric et al, 2010b). Upon

illumination of the sample in the presence of DCMU, a redox pre-steady-state

of the P700 is attained upon oxidation of the PSI donors, after less than one

second. However, if the light is kept on, the fraction of reduced PSI slightly

increases, until a steady-state is achieved. Here it is shown in a figure

adapted from Alric et al (2010), where it is easily visible due to a weak actinic

light:

161

Fig. 28. Typical traces of P700 oxidation in Chlamydomonas in the presence

of DCMU. The curves are a result of substraction of absorption kinetics at 730

nm (PC contribution) from 705 nm changes (PSI and PS redox changes

contribution). Light intensity – 50 e-/s/PSI. Illumination duration 2 or 10 s.

Adapted from Alric, 2010. Top, whole scan. Bottom, zoom on the dark-to-light

or light-to-dark transitions.

In the first panel, a typical trace of PSI oxidation is shown. In the sample

illuminated for 10 seconds (black), there is an apparent reduction of PSI in the

light starting after 3 seconds of illumination. This process influences the rate

of post-illumination PSI reduction rate, as shown in the second panel, and

therefore can be interpreted as an increase in a cyclic process.

We have independently seen this in the same manner:

162

Fig. 29. Typical traces of P700 oxidation in Chlamydomonas in the presence

of DCMU. The curves are a result of subtraction of absorption kinetics at 730

nm (PC contribution) from 705 nm changes (PSI and PS redox changes

contribution). Light intensity – 200 e-/s/PSI. Duration of illumination 1,6, 2,5 or

5 s. Inset, zoom on the light-to-dark transition as marked by the green box.

The increase in the rate of PSI reduction in darkness can be then quantified

by plotting the rate as a function of illumination length:

Fig. 30. Quantification of the rate of post-illumination P700 reduction in the

presence of DCMU. The rate of reduction is plotted against the duration of the

illumination. Mean and standard deviation are shown for three technical

replicates.

163

One can see that this process is independent of the light intensity used for the

illumination, and that the increase in the rate of PSI reduction during the

illumination shows an acceleration of about 1,5e-/s2/PSI.

However, for the reasons outlined throughout the introduction, although we

are in a presence of DCMU one cannot completely exclude that some back-

reactions contribute to the reduction shown here. It is interesting to see

whether the same process exists if probed using the DIRK approach. The

results of this investigation are shown below:

Fig. 31. Results of the multiple DIRK experiment for dark-adapted WT,

dPTOX2, dpgrl1. Each value of the electron flow through the PSI during a 5

second illumination is represented as a point. i values (moments when the

PSI ETR was measured) shown here: 700 1500, 2000, 3000, ms. Each value

is an average of 40 technical replicates each of 2 biological samples.

It is visible that for every strain, the photochemical rate of the PSI increases

during the light period and that the absolute values determined by this

approach and P700 redox measurements are the same. Moreover, as the

light intensities differ vastly between those experiments (30-200 e-/s/PSI for

the P700, 450 e/s/PSI for the DIRK), while the PSI rate does not change, it

substantiates our proposition that in the steady state where the chain is

oxidized, such measurements cannot reveal the actual maximal rate of CEF

as this process is limited by the presence of the substrate – electrons – and

164

that their entry to the membrane is likely kinetically limited by the NDA2 at 2 e-

/s/PSI.

The DIRK-based measurements can also be quantified in order to yield the

rate of increase of the PSI rate in the light:

Fig. 32. Quantification of the increase in the PSI ETR in the presence of

DCMU under illumination. The rate of PSI is plotted as a function of the

moment of probing and each point is an average of 40 technical replicates

each of 2 biological samples. Linear fits for the curves are shown and their

average is reported.

The ‘acceleration’ of the PSI ETR is very similar in this case, with a value of

2e/s2/PSI in these dark-adapted strains. Although this is only an assumption,

we tend to think that it is the 𝑃𝐶𝐸𝐹 that explains changes in the PSI ETR. This

is because there is no reason to expect a significant difference for the NDA2

activity, either by a regulation (modifications of the activity of this enzyme are

unknown) or through the NADPH availability. We therefore hypothesize that

an unknown process modifies the 𝑃𝐶𝐸𝐹.

How can this observation be used to compare it with the transitory CEF? The

𝑃𝐶𝐸𝐹 increase occurs during the first illumination of the dark-adapted sample.

If the decrease, however, back to its dark-adapted level is a long process, the

enhanced 𝑃𝐶𝐸𝐹 probability will be maintained for the second illumination if it

comes fast enough, and will decrease to the basal level if the period of

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darkness is long enough, changing the rate of CEF at the end of the

illumination. We have checked whether it was the case:

Fig. 33. Results of the multiple DIRK experiment Each value of the electron

flow through the PSI during a 5 second illumination is represented as a

column. i values (moments when the PSI ETR was measured): 10, 20, 30, 50,

75, 150, 300, 450, 1000, 2000, 3000, 4000, 5000 ms. dt (time in darkness

after a first oxidation) values: 1, 5, 9, 15, 35 s and dark adapted (shown at 60

s). Each value is an average of 8 technical replicates each for 2 biological

replicates, apart from the dark-adapted value which is an average of 40

technical replicates each of 2 biological samples. Highlighted is the area

corresponding to a (quasi) steady-state CEF to point how does its average

values change depending on the dt.

It can be seen that the rate of CEF at the end of the light period, i.e. when

P700 is mostly oxidized, is higher for the lower dt (time between illumination)

than for the longer dt. We re-plotted the average rate of CEF between 1,2 to 5

seconds of illumination for the WT and dPTOX2 as a function of the dt:

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Fig. 34. Quantification of the average steady-state CEF rate from DIRK

measurements in the presence of DCMU in the WT and PTOX2 mutant. The

average residual rate of PSI (between 1 and 5 seconds of illumination) is

plotted against the time between two illuminations. Exponential fits are shown

for both strains.

As shown above, although arguably the scatter is relatively high (as the PSI

ETR is small in the presence of DCMU), for both strains the CEF is higher

shortly after the first illumination than when the time in darkness becomes

longer.

Hence, we can use these changing values of the ‘residual’ CEF to try to

model the behavior of the transitory CEF. Our reasoning here is the following:

- The increase in CEF at the end of illumination is due to 𝑃𝐶𝐸𝐹

alterations and not NAD(P)H concentration changes.

- The 𝑃𝐶𝐸𝐹 during the transition periods reflects the 𝑃𝐶𝐸𝐹 at the end of

illumination for a given dt.

- The area below the DIRK measurements for various dt is a product

of the quantity of electrons in the membrane before illumination (as

evidenced by the MV measurements) and the partitioning of 𝑃𝐶𝐸𝐹

and 𝑃𝐿𝐸𝐹.

167

We can hence multiply the quantity of electrons in the membrane (P700

DCMU+MV measurements) times the fitted CEF rates from the figure above

(dashed lines), which reflect the changes of 𝑃𝐶𝐸𝐹 over time. We can then

compare the resulting curves with the area under DIRK measurements as

shown before. The result is presented in the following figure:

Fig. 35. Modeling of the measurements reveals that the situation can be

described using only two variables, the starting electrons quantity and the

probability of cyclic. Values calculated from [(number of PSI donors in the

membrane as deduced from the integration of the area under P700 oxidation

in the presence of DCMU and MV) times (normalized CEF efficiency derived

from the fitted function of the residual CEF from the multiple DIRK

measurements)] are reported in red (WT) and green (PTOX2) as a function

of time between illuminations. Evolution of the number of PSI turnovers is

shown for the WT and PTOX2 as calculated from the integrals of multiple

DIRK measurements in blue and orange, respectively, for the sake of

comparison.

We can see that this simple two-component model accounts well for the

explanation of the behavior of the area below PSI ETR measurements. Apart

from the experimental scatter and the initial points during which the pool of

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electron donors was not in steady-state, the correlation of the kinetics is

perfect. Although again this is not a proof that the 𝑃𝐶𝐸𝐹 and our theory is

indeed true, another correlation strongly points in favor of its validation.

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Applications

How can our methods be of use? Assuming we really explore the true

maximal rate of cyclic electron flow with our combined P700-DIRK

measurements, it is of huge interest to use the range of Chlamydomonas

mutants in order to further elucidate the mechanism of this elusive process.

We first wanted to validate our approach with the use of PTOX2 mutant. We

expected two things from the experiments concerning this strain – that the

initial pool of electrons will be larger, because the PQ pool is more reduced in

the absence of the oxidase, and perhaps that a slight increase in the 𝑃𝐶𝐸𝐹

occurs as the stroma of the chloroplast is more reduced in darkness. The

results of the DIRK measurements for the dark-adapted WT and DPTOX2 are

presented below:

Fig. 36. Result of the multiple DIRK experiment for a dark-adapted WT strain

and for the dPTOX2. Each point is an average of 2 biological replicates, each

measured 40 times for the WT, and 4 biological replicates measured 40 times

for PTOX2.

The DIRK measurements comparison between the WT and the mutant yields

following results:

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- The PTOX2 mutant has a higher PSI antenna size. This is expected

because in the dark, the lack of PTOX results in an overreduction of

the PQ pool and migration of the LHCII to the PSI.

- The initial rates of the PSI ETR are similar in both cases, with those

of the mutant being slightly higher.

- After around ~80 ms of illumination, the values of the ETR between

the strains diverge, with higher rates sustained by the mutant. We

interpret this divergence as a start of the oxidation of the PQ pool,

significantly more reduced than in the WT. This at a time increases

the duration of the “linear” electron flow during our experiment

because of a higher initial number of electrons, which also sustains

a longer CEF.

- After a steady-state is achieved, the rates of PSI ETR are slightly

higher with regards to those in the WT, however they bear a

considerable uncertainity (i.e. the measurement error is big when

the slopes differ little).

We can see that the PTOX2 is a great tool for validating our method. With an

access to a bigger pool of PSI electron donors, the behavior of the strain is

the same as the WT, bar the length of high rates sustained (the presence

of the ‘bump’). Importantly, an analysis of the adjusted 𝑃𝐶𝐸𝐹 indicates that it is

the same at the onset of illumination as in the steady-state, as well as being

close to the WT values. Even if the latter is slightly higher (translating into a

modest increase in in CEF rate), the change is not overwhelming. Compared

to our other observations that the NAD(P)H pool is more reduced in the

mutant it could readily explain this difference yet put us in contradiction with

Takahashi 2013 because the rate of CEF reported there was identical to the

WT when corrected by the antenna size. However, the fact that even anoxia-

mediated complete over-reduction of the chloroplast changes the 𝑃𝐶𝐸𝐹

relatively slightly (20-30 pp), points into the direction that a rather large

NAD(P)+ reduction is needed to alter the 𝑃𝐶𝐸𝐹 and perhaps a measurement

error is responsible for the CEF differences in these strains.

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Is is possible to show differences in CEF in mutants affected in this process?

There are few known mutants of cyclic, essentially restrained to pgr5 (proton

gradient regulation 5) and pgrl1 (pgr5-like 1). Both of them were discovered in

Arabidopsis and exist in Chlamydomonas, and are implied in the regulation of

FQR-mediated CEF. Although the mainstream view in the photosynthetic

community is that they “mediate” a “pgrl1/pgr5-dependent” CEF, it has been

shown for pgr5 that a plant mutant lacking this gene exhibits a maximal rate of

CEF that is unchanged compared to the WT. Moreover, the functional

characteristics of the pgrl1 mutant in higher plants are identical to those of

pgr5 (P. Joliot, personal communication), also contradicting the proposed role

of FQR for the pgrl1 (Hertle et al, 2013). We have chosen to employ our DIRK

and P700 measurements in the latter to test our method and hopefully to

elucidate whether pgrl1 regulates or mediates CEF in Chlamydomonas.

Fig. 37. Comparison between the evolution of the number of PSI turnovers

before complete oxidation as measured with area-under-P700-oxidation-curve

methods in the presence of DCMU in the WT or dpgrl1 strains. A first

illumination was applied to oxidize the chain, and a variable time in darkness,

plotted on the x-axis, followed. The area under the lag of PSI oxidation upon

second illumination was divided by a corresponding area of a PC-lacking

mutant, yielding a result in PSI turnovers for the WT (black) and the dpgrl1

(blue). Light intensity – 450 e-/s/PSI in both experiments.

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We first looked at the P700 oxidation. Similarly to previous figures, the areas

under the PSI oxidation traces, normalized to yield a number of apparent PSI

turnovers, were plotted as a function of time spent in darkness between two

illuminations. The initial rise of this value is identical in both strains suggesting

that the rate of PQ reduction during the dark period is the same in dpgrl1 and

in the WT. However, the maximal value reached for the pgrl1-lacking mutant

is significantly smaller than in the WT. As we know, this result can suggest

three things: lower initial number of PSI electron donors, smaller 𝑃𝐶𝐸𝐹 or

maximal CEF rate, or lower reducing pressure at the acceptor side of PSI.

Repeating these measurements in the presence of MV can test the first

hypothesis:

We can see that number of electrons transferred in a LEF-only setup is similar

to that in the WT. We thus decided to use the DIRK-based measurements to

probe whether the decrease of the amplitude of the P700 measurements was

due to CEF differences or to an altered redox status of the stroma.

Fig. 38. Result of the multiple DIRK experiment for a dark-adapted WT strain,

dpgrl1 strain and for the PTOX2. Each point is an average of 2 biological

replicates, each measured 40 times for the WT and pgrl1, and 4 biological

replicates measured 40 times for PTOX2.

The pgrl1 DIRK results are overlayed on the WT and PTOX2 data. One can

see that expectedly, the PSI antenna size is identical (first measurement point

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– the dark-to-light transition) with regards to the WT. However, the rates of

PSI-mediated LEF+CEF diverge quite drastically from the WT values since

the early moments, being in every case lower until 150 ms, where differences

are no longer seen.

We have calculated the maximal rate of CEF according to our theory and

using 𝑃𝐶𝐸𝐹 from the steady-state CEF, similarily to the analysis presented for

the WT. Surprisingly, because of the lower integral the maximal rate of CEF in

the pgrl1 strain is restricted to ~35 e-/s/PSI, a value consistent with maximal

rates of CEF in anoxia in this strain, which were not exceeding 25 e-/s/PSI

(tolleter plant cell), probably again underestimated due to the method

employed. We insist here to say that if true, this is the first demonstration of

the operational pgrl1-‘influenced’ pathway in oxic conditions in

Chlamydomonas.

Lastly, a fourth mutant was tested. This time, although initially by mistake, we

have looked at the oxidation of PSI in an FtsH mutant. FtsH is a thylakoid

ATP-dependent metalloprotease with structural resemblance to some

chaperones (Adam, 2000; Adam & Clarke, 2002; Malnoe et al, 2014). The

mutant we have used lacks the ATP hydrolysis ability due to a point mutation

in the active center of the enzyme. Using the standard approach measuring

CEF in the steady-state, it was shown to have CEF rates slightly higher than

the WT (A. Malnoe, PhD thesis). The results of the P700 oxidation

experiments we have conducted are however extremely interesting because

the mutant exhibits a completely different kinetics than the previous strains:

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Fig. 39. Comparison between the evolution of the number of PSI turnovers

before complete oxidation as measured with area-under-P700-oxidation-curve

methods in the presence of DCMU in the WT, pgrl1, and various FtsH

strains. A first illumination was applied to oxidize the chain, and a variable

time in darkness, plotted on the x-axis, followed. The area under the lag of

PSI oxidation upon second illumination was divided by a corresponding area

of a PC-lacking mutant, yielding a result in PSI turnovers for the WT (black),

dpgrl1 (blue), or FtsH (light blue, green, red). Light intensity – 450 e-/s/PSI in

each experiment.

It is visible that the initial rate of the reduction is faster, the amplitude is

smaller, and the rate of oxidation faster than in the WT; moreover, the dark-

adapted strain (dt=60 s) shows an extremely limited number of turnovers

before PSI oxidation, which is not due to an oxidized donor pool. The latter

was tested with MV (not shown) and it turns out that the pool is even more

reduced than in the WT. Hence, the mutant does not cycle nearly any of the

electrons upon illumination, suggesting a vast alteration of the CEF

mechanism. This observation is even more striking when taken into

consideration that:

- FtsH is found to co-migrate with auxiliary CEF proteins on sucrose

density gradients (F. Buechert & F.-A. Wollman, personal

175

communication) in what resembles a ‘CEF supercomplex’ (Iwai et

al, 2010a).

- FtsH co-precipitates with pgrl1 (F. Wang, C. de Vitry, M. Hippler

and F.-A. Wollman, personal communication), suggesting a direct

interaction between these proteins.

Further steps need to be undertaken in order to fully understand the role of

CEF-related proteins, however we feel that our method can be an important

tool for this.

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State transitions

Introduction During the course of my PhD, nearly simultaneously in early 2014, two

research articles about state transitions in Chlamydomonas were published in

PNAS. The first one, by the groups of van Amerongen and Croce examined

the process using fluorescence lifetime analysis in vivo. The second one, a

joint effort from Minagawa, Finazzi and Garab laboratories analyzed changes

in thylakoid membrane structure upon state transitions using a variety of

approaches (Nagy et al, 2014).

These publications can be considered a follow-up of a previous paper from

Minagawa’s group (Iwai et al, 2010b), who also used fluorescence lifetime

imaging to explore state transitions. Finally, after these initial articles, Dekker

studied fluorescence emission from state I and state II cells in 77K

(Wlodarczyk et al, 2015).

The common denominator of these papers is the following: the authors

postulate that upon state transitions a significant fraction of the mobile

antenna, LHCII which detaches from photosystem II upon transition from

State I to State II, does not migrate to the non-appressed regions of the

thylakoid membranes where it would increase the functional size of PSI.

Instead, this fraction of the mobile antenna forms aggregates that efficiently

quench the excitation energy. The authors thus suggest that whereas state

transitions upon unbalanced excitation energy between the two photosystems

essentially are identical in plants and in algae, the algae-specific transition

from oxic to anoxic conditions involves an additional process, in which a major

decrease of PSII antenna size occurs with no parallel increase in PSI antenna

size.

Interestingly, proposals that State transitions do not increase PSI cross-

section date back. As pointed out by Allen (1992), there was a debate in the

literature in the eighties whether there is an actual increase of the PSI

antenna size concomitant with FMax decrease (due to PSII antenna

detachment). The few attempts to measure the rate of P700 oxidation were

unsuccessful in detecting such PSI antenna changes and concluded similarly

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to the recent findings that it remained below detection (Allen & Melis, 1988;

Haworth & Melis, 1983). However, in Chlamydomonas it has been accepted

that state transitions consist of a massive decrease of the PSII antenna size

(50%) and a concomitant increase in PSI absorption cross-section. This was

demonstrated in our laboratory in 1985 and 1996 (Delepelaire & Wollman,

1985; Delosme et al, 1996).

The implications of these findings are of major significance in the field of

photosynthesis since state transitions would no longer provide a balancing

mechanism, where a decrease of quinone-reducing PSII activity would be

further enhanced by an increase of PQH2 oxidation, since it would serve only

a quenching purpose. From the evolutionary standpoint, it would question the

rationale of retaining state transitions as a major regulation of the

photosynthetic apparatus in green algae instead of qE as in higher plants (in

Chlamydomonas the development of qE requires induction in high light and

photoautotrophic conditions). The latter has several advantages over state

transitions as its response is more rapid, the efficiency of quenching greater, it

does not necessitate displacement of any proteins in the membrane and is

ATP-independent.

The view above also is poorly consistent with the ecological niche of green

algae. Growing in soil or in shallow ponds these organisms are often exposed

to hypoxic conditions and it is known that their metabolism, gene expression

and photosynthetic parameters vastly change upon transition to anoxia (for a

comprehensive review, see (Yang et al, 2015)). In these conditions light is

much less abundant than for higher plants and PSI needs to cope not only

with PSII activity, but also with the light-independent quinone reduction via

NDA2, the substrate of which, NAD(P)H, is abundant following deprivation in

O2, the terminal electron acceptor of mitorespiration. For this reason, an

increase in photosystem I antenna size is theoretically even more crucial then

the downregulation of PSII.

In view of these rather intriguing new suggestions regarding state transitions

that, in addition, were fully conflicting with numerous previous studies from our

laboratory (Delepelaire & Wollman, 1985; Delosme et al, 1996; Vallon et al,

178

1991), we decided to reassess the changes in the respective antenna sizes of

each photosystem upon state transitions using time-resolved spectroscopy in

vivo. We reasoned that by addressing these questions in whole cells with

unambiguous methods such as by probing the rate of P700 oxidation in the

light we would avoid short-comings from biochemical preparations (Nagy et al,

2014) and indirect interpretation as those arising from deconvolution of

fluorescence lifetime signals (Iwai et al, 2010b; Unlu et al, 2014). We thought

that working in vivo at room temperature is the most reliable approach

because fragile protein-protein interactions or thylakoid structure could be

affected at low temperature (Wlodarczyk et al, 2015). We thus relied on 77K

emission spectra only for a qualitative crosscheck of our conditions. We also

wanted to place emphasis on the reproducibility of the experiments – because

the signals measured are rather small and their variations even less

pronounced. Thus, it was crucial to establish a proper statistical view of state

transitions.

179

State transitions paper

180

Topic: Photosynthesis

State transitions redistribute rather than dissipate energy

between the two photosystems

Short title: State Transitions redistribute excitation influx

Wojciech J. Nawrocki1#, Stefano Santabarbara2#, Laura Mosebach1,3, Francis-

André Wollman1*, Fabrice Rappaport1*

1: Institut de Biologie Physico-Chimique, UMR 7141 CNRS-UPMC, 13 rue P.

et M. Curie 75005, Paris, France

2 : Istituto di Biofisica, Consiglio Nazionale delle Ricerche, Via Celoria 26,

20133 Milano, Italy.

3: present address: Munster University, Institute of Plant Biology &

Biotechnology, D-48143 Muenster, Germany

#: both authors contributed equally to the work.

*: Corresponding authors: fabrice.rappaport@ibpc.fr; francis-

andre.wollman@ibpc.fr

181

Abstract:

Photosynthesis feeds the biosphere in energy by converting sunlight into

biologically useful compounds. This catalytic feat involves two photosystems

acting in series and powered by Light Harvesting Complexes which increase

the flux they sustain by many folds. These complexes are the main targets of

the regulatory processes that allow photosynthetic organisms to cope with a

broad range of light intensities. In microalgae, which attract considerable

biotechnological interest owing to the potentialities offered by their metabolic

plasticity, one of such processes, State Transitions have much larger

amplitude than in terrestrial plants, whereas thermal dissipation of energy, the

dominant regulatory mechanism in plants, only takes place after acclimation

to high light, suggesting distinct and complementary roles for these two

mechanisms. Here we show that, at variance with recent reports, in

microalgae, State Transitions do not dissipate light energy but redistribute it

between the two photosystems, thereby allowing a well-balanced influx of

exciting energy.

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Introduction:

Oxygenic photosynthesis requires two photosystems, Photosystem II (PSII)

and I (PSI), acting in series to catalyze the transfer of electrons from water to

CO2. These two membrane-bound complexes convert light energy, via

electron and proton transfer reactions, into electrochemical energy and

ultimately into biologically-useful free energy1. But before being converted,

light energy must be captured and the catalytic feat that these complexes

achieve would be limited without the light harvesting complexes (LHC) that

increase the flux they sustained by more than two orders of magnitude

funnelling light energy to the photochemical converters and thereby

enhancing their absorption cross section (reviewed in 2).

Unsurprisingly, light harvesting is the major target of regulatory processes of

photosynthesis3. In high light conditions, when the excitation flux exceeds the

assimilation capacity, the limiting step of photosynthesis is one of the light

independent electron transfer reactions, and regulatory mechanisms are

triggered that decrease the excitation pressure and the resulting over-

reduction of the electron acceptors, a potent source of light-induced damages.

Prominent among these is Non-Photochemical Quenching (qE), which

increases the thermal dissipation of the excited state of

chlorophylls4,_ENREF_5.

In low light conditions, the turnover rate of each photosystem is proportional to

the energy influx it receives and the least excited photosystem sets the

velocity of the entire chain. Consequently, whereas a balanced excitation flux

warrants a smooth operation in a dynamic range of light intensities, an

unbalanced excitation influx generates a suboptimal oxidation state of the

electron carriers in the chain and compromises its ability to cope with large

and sudden changes in light intensity. This becomes critical in microalgae

often exposed to hypoxic environmental conditions under which the

plastoquinone (PQ) pool that shuttles electrons between the two

photosystems becomes over-reduced5.

Since their discovery in microalgae in the late 1960’s, “State Transitions“ (ST)

have been regarded as responsible for preserving a balanced excitation

between the two photosystems6,7 (reviewed in8,9). ST involves the reversible

phosphorylation of LHC antenna proteins and their consecutive lateral

183

migration from the PSII-enriched stacked membrane regions of the thylakoid,

to the PSI-enriched unstacked ones10,11. The reversibility of this redistribution

relies on the interplay of a phosphatase and a kinase that is activated by the

reduction of the PQ pool sensed by the cyt. b6f complex_ENREF_12

(reviewed in8). Thus, in unicellular green algae, ST would adjust the relative

antenna sizes of the two photosystems using the redox state of PQ’s as a

sensor of their uneven light excitation or of changes in light-independent

metabolic processes that impact the redox state of the chloroplast stroma

and, consecutively, of the photosynthetic electron transfer chain. In this view,

the respective light harvesting capacity of each photosystem would change in

a compensatory manner with the mobile LHC commuting between the two

photosystems, not only to balance the excitonic flux but also to meet the cell’s

demand for ATP upon changes in environmental conditions12.

Yet, this view has been repeatedly challenged in the 1980’s13 and more

recently in studies 14-17 that concluded that microalgae undergo ST of much

larger amplitude than vascular plants because a major fraction of the mobile

antenna detached from PSII remains disconnected from PSI and aggregated

in a quenched state. Accordingly, green algae such as Chlamydomonas

reinhardtii would be equipped with only one major regulatory mechanism

dissipating energy, through both ST and qE under low and high light

respectively, while there would be only a minor contribution of ST in balancing

the excitation flux under low light conditions.

This is however hard to reconcile with the physiology of Chlamydomonas

found on soil and in shallow ponds, where light intensities are rarely excessive

but that are prone to generate hypoxic conditions, as documented by a

metabolic flexibility that plants do not develop18. This idiosyncratic feature

calls for a preserved photosynthetic activity in order to keep high levels of

intracellular ATP when the PQ pool is reduced in the dark in response to

hypoxic/anoxic conditions and it also rationalizes the inability of

Chlamydomonas to develop a qE-type of quenching unless acclimated to high

light beforehand19.

The paradoxical conclusions conflicting with the ecology of green microalgae,

such as Chlamydomonas, which have attracted huge biotechnological interest

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owing to the potential of their metabolic plasticity, prompted us to reassess

the mechanism of state transitions in this unicellular model organism.

Results:

We first sought a direct method to measure the PSI antenna size and its

putative relative change upon State Transitions. We reasoned that measuring

the oxidation rate of P700 in a mutant lacking plastocyanin meets this

purpose since, in such a mutant, each charge separation generates a long-

lived P700+

due to the absence of a secondary donor. Thus the rate of

formation of P700+ in continuous light is determined by the rate at which PSI is

excited which is proportional to the antenna size and to the light intensity.

Such a mutant however prevents the tight control of the redox state of the PQ

pool and thus the achievement of a complete State I. Indeed, in the absence

of PC, the PQ pool cannot be fully oxidized by illumination and inhibition of

PSII, owing to the interruption of the chain upstream of PSI. State I conditions

were thus promoted by a vigorous aeration of the cells in darkness by

shaking, a method that has been reported previously to produce up to 80% of

the completion of State I 20,21. State II was achieved by placing the cells in

anoxia, as in15,16.

Figure 1 shows the kinetics of the light-induced absorption changes at 705 nm

with increasing light intensities in both State I and State II conditions. As

shown in the inset, the oxidation rate of P700, determined by fitting the

oxidation kinetics with one exponential, increased linearly with the light

intensity. Saliently, the slope of this linear plot increased by 39% ± 10%

(n=13) upon a shift from State I to State II (inset of Figure 1 and Figure 2). We

note that, whereas a single exponential yielded an excellent fit in State II,

using two exponentials increased the quality in State I (see figure S1).

Interestingly, according to the bi-exponential fit, in State I the fast component

accounted for ~20% of the total amplitude and was 1.4 fold faster than the

slower component, which, in its turn, had a similar rate as that found in State

II. This suggests that, in the State I conditions we used with this mutant, ~20%

of the mobile LHC is already bound to PSI in a State II configuration.

We next examined how the relative weight of the two components contributing

to the P700 oxidation kinetics evolved as State Transition took place. We

globally fitted the series of individual kinetics measured during the time course

185

of ST with a sum of two exponentials. We found that the relative amplitude of

the fastest component progressively increased at the expense of the other

one within 3 minutes after the induction of the ST (Figure 3), consistent with

previous reports on the rate of ST22. Accordingly, ST would involve a transient

mixture of two types of PSI, having respectively a “State I like” and a “State II

like” antenna, rather than the progressive increase of the absorption cross

section of each individual PSI.

We then asked how these changes in PSI antenna size compare with the

simultaneous variations in PSII antenna size. These variations are commonly

assessed by measuring the change in fluorescence intensity under conditions

where PSII photochemical traps are “closed” (FMax), i.e. unable to generate a

stable radical pair20. At FMax, the emission is dominated by PSII and its

intensity is proportional to the probability that a photon is absorbed by PSII.

We thus measured in parallel to the P700 oxidation kinetics, i.e. with the same

sample and in the same conditions, the decrease of FMax associated with the

transition to State II in the mutant. We found a decrease of 35% ± 5%

(Figure 2) which is similar to the 39% ± 10% increase estimated above for the

change in the relative antenna size of PSI. However, to assess the

complementarity of these changes in antenna, one needs to know the

antenna size of PSII relative to that of PSI before the transition takes place.

We thus assessed the relative cross section of PSII and PSI using the

Electrochromic shift (ECS) undergone by carotenoids and chlorophylls b

bound to LHC proteins23. The absorption changes resulting from this shift are

proportional to the transmembrane electric field generated by photochemical

charge separations. The ECS signal induced by a saturating and single

turnover flash is thus proportional to the total number of charge separations

undergone by all photosystems (I and II). Upon inhibition of PSII by specific

inhibitors, the contribution of PSI alone and, after subtraction, that of PSII, can

be determined. We thereby found that the PSII to PSI ratio per electron

transfer chain was ~1 (Table 1). When using a weak, sub-saturating exciting

flash, the probability that a given photosystem undergoes charge separation is

determined by the probability that it traps an absorbed photon. Thus, when

measured under non-saturating conditions, the ECS signal associated with

186

PS I and PSII is proportional to their respective antenna size. In the

mutant, we found the ratio between the PSI and PSII absorption cross section

to be 1.2 ± 0.1 (n=7) in State I (Table 1), which is consistent with the relative

amplitude of the fast component in the P700 oxidation kinetics in State I (Figure

3). It also is in perfect agreement with previous photoacoustic measurements

with a dark-adapted WT strain20.

When assessing the change in the relative absorption cross section of PSI

upon State Transition using ECS, we found that it increased by 28% ± 5%

(see Figure 2). Although far from being negligible, this change is smaller than

that estimated by the direct measure of the P700 oxidation rate. This apparent

discrepancy between the ECS and P700 oxidation methods is readily explained

by our above observations that a fraction of phosphorylated LHCII is already

bound to PSI in State I conditions in the mutant. Indeed, while the latter

method leads to the complete oxidation of P700 and thus probes the entire

population of PSI, the former method, relying on sub-saturating single

turnover flashes, only probes a fraction of PSI centres. In State I this fraction

will be enriched in those PSI centres having the larger absorption cross

section and the method will overestimate the antenna size of PSI in State I, in

turn underestimating the cross section increase upon transition to State II.

We then studied ST in the wild type strain, grown in photoautotrophic

conditions. Complete State I was achieved by fully oxidizing the PQ pool via

illumination in the presence of DCMU to inhibit PSII. State II was promoted as

for the mutant, by anoxia. The more pronounced State I achieved with

the wild type relative to that in the mutant resulted in larger changes in

77K emission spectra, a method which is routinely used to asses State

Transitions24 (see figure S2).

Using the ECS method described above we found that the relative absorption

cross section of PSI with respect to PSII in State I, was of 1.0 ± 0.1 (n=9;

Table 1). With the same method, we found a 35% ± 9% (n=14) increase of the

PSI absorption cross section when ST were completed from State I to State II.

This is to be compared to a decrease by 38% ± 6% (n=14) of PSII antenna

size as assessed by the changes at FMax. Given this remarkable

complementarity of the changes in antenna size of the two photosystems in

the wild type strain of Chlamydomonas, we conclude that virtually all of the

187

peripheral antenna that detaches from PSII upon transition to State II,

functionally couples to PSI.

The present data fully support a redistribution of the light harvesting antenna

between the two photosystems upon ST in Chlamydomonas, but conflict with

any significant contribution of the aggregation-quenching LHCII model that

has been recently proposed. To further re-assess the two models, we

revisited the situation in a mutant lacking the PSI reaction centres. This

mutant is predicted to have opposite behaviours upon transition to State II

depending on the model one considers. The “antenna redistribution model”

would predict no changes of FMax, because of the absence of the quenching

sites provided by PSI. The “aggregation-quenching model” would predict the

same extent of decrease at FMax as in the WT, the quenching being

independent from PSI. In agreement with the “antenna redistribution model”

we observed no quenching at FMax in this mutant upon transition to State II, in

line with Delosme et al.20. We checked that the unchanged FMax was not due

to the lack of phosphorylation of the antenna protein that expectedly displayed

the characteristics of State II in the ΔPSI mutant (see Supplementary Figure

S3). Thus PSI must be present to promote the quenching of fluorescence in

State II, consistent with earlier conclusions that the absence of PSI in

Chlamydomonas20 or the lack of specific PSI subunits in PSI of A. thaliana25

impeded ST.

Discussion:

In unicellular green algae, State Transitions (Figure 4) have long been

considered as a mechanism balancing the excitation flux of PSII and PSI by

the shuttling of a mobile fraction of the antenna. However, this view has been

repeatedly challenged (see 13). Recent reports 14-17 concluded to unique

features of ST in the microalga Chlamydomonas that would have developed

self-quenching properties of its antenna proteins when placed in State II

conditions. Contrarily to these statements, we demonstrated with the present

data that ST induce complementary variations of the antenna size of the two

photosystems in Chlamydomonas and thus rebut the conclusions that a major

fraction of the phosphorylated LHCII detached from PSII does not attach to

PSI.

188

We resorted to a plastocyanin lacking mutant that allows the direct measure

of the PSI antenna size as well as of the homogeneity of the absorption cross

section of the PSI population. This mutant, despite having the drawback of not

allowing achieving a full State I, also provided a picture of the dynamics of the

transient homogeneity/heterogeneity of the PSI antenna during ST. We

observed that the absorption cross section of PSI increases significantly upon

transition to State II, and that this increase is commensurate to the decrease

of the PSII antenna. An alternative approach relying on ECS measurements

provided additional support by evidencing, in the mutant and in the WT

strain grown under photoautotrophic conditions, the complementary

redistribution of the respective antenna of the two Photosystems by 35-40%.

Last, the absence of fluorescence quenching at FMax in State II conditions

when using a PSI-lacking mutant showed that PSI is required to trap the

excitation of phospho-LHCII detached from PSII.

As regards to the assessment of the PSII antenna from FMax data at room

temperature, a reliable determination of the variation in number of chlorophylls

connected to PSII requires taking into account the small but not completely

negligible contribution of the PSI photochemical units to the overall

fluorescence emission. It has been estimated as 15-30% of F026 which, in C.

reinhardtii that has typically an F0/FMax ratio of 0.3, leads to a contribution of

PSI to FMax of ~5-10%, at the relevant detection wavelength27. Thus the

variations of FMax by 35% and 39% in the and WT strains correspond to

changes in the PSII antenna size of 37-39% and 41-43%, respectively.

But is the present finding that about 35-40% of antenna moves between PSII

and PSI upon ST quantitatively compatible with the current knowledge on the

biochemical and pigment contents of this mobile antenna? These data,

available from the literature, are recapped in Table 2.

According to literature, in C. reinhardtii, PSII bears 190-230 (we’ll thereafter

use 210 as a conservative figure) chlorophylls in State I and PSI 220 (see

Table 2) Consistent with this, we found that PSI and PSII have similar

antenna size in the WT in State I. In the mutant, the antenna size of PSI

relative to PSII was 1.2 and this is explained by the incomplete oxidation of

the plastoquinone pool in the experimental conditions used to generate State

I. Therefore, assuming no disconnected chlorophylls and neglecting the

189

slightly smaller trapping efficiency of PSII versus PSI (because, considering

the intrinsic scatter of biological data, this would be a second order

correction), the antenna size would be 195 Chls for PSII and 235 Chls for PSI

in State I in this strain. With this in mind and considering the biological and

experimental uncertainties of these figures, the 38 % decrease of the PSII

antenna size in the ∆PC mutant would involve the disconnection of ~75 Chls

from PSII that, when bound to PSI would result in a 33% increase of its

absorption cross section, in reasonable agreement with the 39% found here.

As to the WT, the 42 % decrease in the PSII antenna size corresponds to the

detachment of ~85 Chls, which also matches the 35% complementary

increase of the PSI absorption cross section.

These estimates also fall in line with28,29 who characterized the LHCII-LHCI-

PSI supercomplexes isolated from State II conditions They contained – apart

from the constitutive 9 LHCI and the core – two LHCII trimers and a

monomeric peripheral antenna28 or 5 antenna subunits29. In terms of

chlorophyll per Photosystem, this adds 97 or 70 Chls, respectively, which,

assuming identical efficiency for the energy transfer to the core as from the

LHCI chlorophylls30,31 translates into a 33-46 % increase in the antenna size

compared to State I, in good agreement with our results (39-44%).

Importantly, it means that in complete State II, all the PSI-LHCI

supercomplexes bind at least 2 phospho-LHC trimers.

Owing to biological scatter and experimental errors (see the width of the

Gaussian distribution in Figure 2), we cannot definitely exclude the existence

of a fraction of unbound LHCII in State II conditions but the present data show

that it must be minor. A similar conclusion was also reached by Wlodarczyk et

al. who reported, based on the study of the fluorescence emission lifetime at

77K and 6K, the presence of a yet uncharacterized component emitting at 685

nm, that they assigned to unbound LHCII and which accounted for 20% of the

mobile LHC pool, the remaining 80% being bound to PSI32. According to our

data, obtained at room temperature, the maximum fraction of disconnected

and quenched LHC would be closer to 10% and the possibility remains that

structural modifications induced by freezing of the thylakoid membranes in low

190

temperature measurements lead to an overestimation of the detached LHCII

pool.

Thus, as depicted in Figure 4, we found that in State II, in C. reinhardtii, the

vast majority of PSI centres bind 2 LHCII trimers and probably another

monomeric antenna33. The increase in PSI antenna size agrees well with

biochemical and structural data and so does the complementary decrease in

that of PSII. The extent of ST in microalgae is therefore higher than in

vascular plants, and contrary to those recent reports that suggested an LHCII

aggregation-quenching process14-16 (see supplementary information for

further discussion), it consists mostly of photochemical quenching by PSI. The

antenna redistribution model suits well with the ecological niche the algae

inhabit, Chlamydomonas being often exposed to hypoxic or anoxic conditions,

when growing on soil in dense populations18, conditions that are never met by

land plants This habitat also results in low light exposure, as well as in the

presence of reduced carbon sources from the microbial environment. The

physiology of ST is then to increase the photochemistry of PSI to compete

more efficiently with both photochemical and non-photochemical reduction of

electron carriers – the latter being important in algae due to a different sugar

metabolism in the chloroplast and an increased rate of chlororespiration5,34.

Consistent with this, a Chlamydomonas mutant having impaired respiration,

and thus an over-reduced stroma in the dark, shows impaired growth in light-

limiting conditions when unable to perform ST12. Indeed, the increased PSI

antenna size in State II conditions, combined with the induction of the

hydrogenase, insures an efficient recovery from over-reducing conditions in

hypoxia/anoxia - allowing rapid restoration of linear and cyclic electron flows,

a critical issue for algal survival in its natural habitat35,36.

Materials and methods:

Strains, growth conditions and treatments

Chlamydomonas reinhardtii strains used in this study are wild type and mutant

progeny from a cross between a plastocyanin lacking mutant ac20837 and a

control strain from our laboratory, both derived from the original wild type 137-

C_ENREF_38. These strains therefore are in a similar genetic background38.

Wild-type was grown in photoautotrophic conditions in Minimum medium18

191

under moderate light (30 µE.m-2.s-1). The plastocyanin lacking mutant (∆PC)

was grown in heterotrophic conditions, in TAP medium18 containing acetate

as a reduced carbon source, at 30 µE.m-2.s-1. Cells were harvested in mid-log

phase (1-4 106 cells.mL-1), concentrated ~6 fold in Minimum medium

supplemented with Ficoll 400 (10% w/w), and shaken in darkness in open

flasks at 300 rpm for at least 30 mins prior to measurements. To induce State

II, glucose (20 mM) and glucose oxidase (50 u/mL; type II, from A. niger) were

used. State I conditions were achieved by light-DCMU (10 µM) treatment

(WT) or by vigorous shaking in the dark (∆PC). As a control strain unable to

perform State Transitions we use the stt7-9 strain12 which lacks the kinase

responsible for the phosphorylation promoting the transition from State I to

State II. As a mutant lacking PSI, we used a strain deleted for the psaB gene

which encodes one of the two main subunits of the core of PSI39.

Time resolved absorption spectroscopy.

Light-induced absorption changes were measured as previously described

in40,41, with a JTS 10 (BioLogic, Grenoble) spectrophotometer that uses

pulsed LED as detecting light sources. The appropriate detection wavelengths

were selected using interferences filters: 705 nm, 10 FWHM to measure the

oxidation kinetics of P700, 520 nm, 10 nm FWHM and 546 nm, 10 nm FWHM

to measure the electrochromic shift absorption changes. 640 nm emitting

LED’s were used as continuous exciting light source. Single turnover flashes

were provided by a dye laser (DCM, Exciton dye laser) pumped by a

frequency doubled Nd:YAG Laser (Minilite II, Continuum). Actinic light was

filtered by a 3 mm thick RG695 Schott glass filters when detecting at 705 nm

and 3 mm BG39 Schott filters when detecting at 520 nm and 546 nm. The

ECS absorption changes were determined after subtraction of the absorption

changes measured at 546 nm to those measured at 520 nm.

Since DCMU (10 µM) was used systematically in the WT to achieve a

complete State I, this PSII inhibitor was also added to the strain when

measuring the oxidation of P700 for the sake of consistency. In addition

hydroxylamine (10 mM) was added, when required, to cancel the contribution

of PSII to the ECS signal measured after a single turnover flash. The mono-

and biexponential fitting was done with OriginLab software. To check that the

192

possible contribution of fluorescence changes to the signal measured at 705

nm were negligible, we checked that the P700 oxidation-induced absorption

changes at 820 nm, where the contribution of fluorescence should be smaller,

were in good agreement with those measured at 705 nm (see Supplementary

Figure S4).

Fluorescence spectroscopy

Fluorescence measurements were measured using the JTS 10 (Biologic,

Grenoble) spectrophotometer in ‘fluorescence’ mode as described

previously26. The fluorescence yield was probed using broad band blue

detecting pulses. To warrant homogenous illumination throughout the sample

green light was used as an actinic light (peak emission at 530 nm, FWHM 50

nm). The detecting and actinic light was filtered out using a combination of a

long pass reflecting filter (50% transmission at 650 nm) and an RG665 Schott

filter. The samples were poisoned with 10 µM DCMU prior to the experiments

to maintain the plastoquinone pool oxidized in both oxic and anoxic

conditions.

77 K fluorescence emission spectra

Low temperature emission spectra were measured using a laboratory built

set-up. The samples, prepared as just described above, were placed in an

aluminium sample holder which was plunged into liquid nitrogen. The

fluorescence exciting light was provided by a LED max 470 nm) and shone

onto the sample using a Y-shaped optical fibre. The other branch of this fibre

was connected to a CCD spectrophotometer (QE6500, Ocean Optics) to

measure the emission spectrum.

Isolation of thylakoid membranes, western blotting and

immunodetection with an anti-phosphothreonine antibody were performed

as previously published42.

193

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nine light-harvesting complexes (Lhca) located on one side of the core.

The Journal of biological chemistry 286, 44878-44887,

doi:10.1074/jbc.M111.301101 (2011).

46 Barber, J. REGULATION OF ENERGY-TRANSFER BY CATIONS

AND PROTEIN-PHOSPHORYLATION IN RELATION TO THYLAKOID

MEMBRANE ORGANIZATION. Photosynth. Res. 10, 243-253,

doi:10.1007/bf00118289 (1986).

47 Kargul, J. et al. Light-harvesting complex II protein CP29 binds to

photosystem I of Chlamydomonas reinhardtii under State 2 conditions.

Febs J. 272, 4797-4806, doi:10.1111/j.1742-4658.2005.04894.x

(2005).

48 Suga, M. et al. Native structure of photosystem II at 1.95 A resolution

viewed by femtosecond X-ray pulses. Nature 517, 99-103,

doi:10.1038/nature13991 (2015).

49 Mazor, Y., Borovikova, A. & Nelson, N. The structure of plant

photosystem I super-complex at 2.8 A resolution. eLife 4, e07433,

doi:10.7554/eLife.07433 (2015).

50 Tokutsu, R., Kato, N., Bui, K. H., Ishikawa, T. & Minagawa, J.

Revisiting the Supramolecular Organization of Photosystem II in

199

Chlamydomonas reinhardtii. Journal of Biological Chemistry 287,

31574-31581, doi:10.1074/jbc.M111.331991 (2012).

200

Correspondence and material requests should be addressed to:

fabrice.rappaport@ibpc.fr / francis-andre.wollman@ibpc.fr

Acknowledgments:

We thank Sandrine Bujaldon for the -WT cross. W.J.N., F.-A.W and F.R.

acknowledge the support of CNRS and Université Pierre et Marie Curie. This

work was supported by the Initiative d’Excellence program from the French

State (grant DYNAMO, ANR-11-LABX-0011-01) and by the program PHC

PROCOPE 2014 PROJET N° 30729YB. W.J.N. is a recipient of a PhD

fellowship from Université Pierre et Marie Curie. S.S. acknowledges the

support from the DYNAMO, ANR-11-LABX-0011-01 grant for travel. L.M. was

a recipient of Studienstiftung des Deutschen Volkes fellowship for her Master

studies.

Author contributions:

W.J.N., S.S., L.M performed the experiments. F-A.W, F.R conceived and

designed the research. All of the authors discussed, analyzed the data and

wrote the paper.

Competing interests statement:

The authors declare no competing interests as defined by Nature Publishing

Group, or other interests that might be perceived to influence the results

and/or discussion reported in this article.

201

Table 1. The relative antenna size ± standard deviation of PSI and PSII in the

Table 2. A survey of the stoichiometry of chlorophylls per photosystems and

LHC’s recapping the current knowledge from the literature regarding C.

reinhardtii and the conclusions drawn from the present study. x

Figure 1. Kinetics of P700 C. reinhardtii mutant. The light-

induced oxidation of the P700 dimer of chlorophylls of PSI was recorded at 705

nm. Experimental data are shown as symbols and the solid lines are

monoexponential fits of oxidation kinetics. These were measured at different

light intensities in both State I and II. The inset shows the linear dependence

of the oxidation rate determined by the monoexponential fit on the light

intensity in both State I and II. The ratio between the two slopes yields the

change in the PSI antenna size upon ST (1.4 ± 0.1). The data shown in the

inset are the average of n=13 independent biological replicates.

Figure 2. Comparison of the changes in the PSII and PSI antenna size upon

State I to State II transitions using different approaches as detailed in the text.

The open symbols show the results of individual experiments, the solid

symbols are the average n≥ 13 of biological replicates and the vertical lines

show the standard deviations. The bell-shaped curves show the

corresponding Gaussian distribution to illustrate the statistical scatter and the

consecutive requirement for numerous biological replicates.

Figure 3. Evolution as a function of the time after induction of ST of the fast

and slow components in the oxidation kinetics of P700

sample was initially placed in State I and then anoxia was induced at t=0.

Oxidation kinetics of P700 were measured at discrete time points following the

induction of the transition to State II. They were globally fitted with two

202

exponentials leaving both the amplitude and life time as free running

parameters. The fit yielded two components with respective time constant of

8.7 ± 0.1 ms (blue squares) and 5.9 ± 0.2 ms (orange circles); note that the

ratio between the two =1.4. The time course of the changes of the relative

amplitudes of these two components as ST develop is shown and the overall

time constant of the process was found to be 176 s ± 10 s. Each single point

is the average of 3 biological replicates at one light intensity.

Figure 4. A quantitative scheme of photosystems and antenna distribution

heterogeneity and of its change during State Transitions in Chlamydomonas

reinhardtii. In state I, when the PQ pool is oxidized, PSII-(3/4) LHCII

supercomplexes form semi-crystalline arrays in the stacked regions of the

thylakoids43,44_ENREF_43, whereas PSI form supercomplexes with 9 LHCa

proteins45 mostly in the unstacked regions11. The functional antenna size of

these photosystems is identical when PQ pool is fully oxidized (this article).

Upon reduction of the PQ pool, phosphorylation of PSII core and antenna

occurs, leading to electrostatic repulsion from the stacked regions of the

membrane13,46 and their migration to the unstacked regions10,20, loosening the

overall stacking16. In state II, PSI-9 LHCa supercomplex further increases its

size by about 40% (this article) by attaching two LHCII trimers and a

monomeric antenna such as CP26/CP29 or 5 antenna subunits28,29,33,47.

203

(dark, oxic) WT (DCMU, light, oxic)

PSI:PSII antenna size ratio in State I 1.2 ± 0.1 (n=7) 1.0 ± 0.1 (n=9)

PSI:PSII ratio 0.9 ± 0.1 (n=7) 1.0 ± 0.1 (n=4)

204

Chlamydomonas reinhardtii PSII (monomer) PSI PSI/PSII

antenna size

State I

Core size 35 chls48

97 chls49

1 ± 0.1

(this work)

Antenna pool

size

~3-4 LHCII trimers

= 126-168 chls +

CP26, CP29 = 26

chls44,50

9 LHCI ≈ 126 chls45

Total

size

~190-230

chls ~220 chls

Oxic,

darkness

(incomplete

STI)

Total size ~210 chls ~250 chls

1.2 ± 0.1

(this work;20

)

% of change

(this work)

From STI - 42 ± 6 % + 35 ± 9 % (ECS)

-

From

incomplete

STI

- 38 ± 5 % + 39 ± 10 % (P700

oxidation)

Mobile chls

2LHCII trimers + CP26/CP2928,45

= 97 (46%)

5 antenna subunits29

= 70 (33%)

State II

Core size 35 chls48

97 chls49

2.35

(this work) Antenna pool

size

~2 LHCII trimers =

84 chls

CP26/CP29 = 13

chls (44

; this work)

~9 LHCI ≈ 126 chls45

+

~2 LHCII trimers = 84

chls

CP26/CP29 = 13

chls28

205

or

~2 LhcbM5 + ~2 CP29

+ ~1 CP26 = 70 chls29

Total

size ~135 chls ~300-320 chls

206

Figure 1.

207

Figure 2.

208

Figure 3.

209

Figure 4.

210

Supplementary data

Supplementary Figure S1:

The kinetics of P700 oxidation measured at 705 nm in the PC mutant fitted

using mono- and bi-exponential functions. On the upper panel, the P700

oxidation data in the PC mutant in State I conditions was fitted with either

mono- (blue line) or bi-exponential function (orange line). The lower panel

shows the residuals between the experimental values and corresponding fits

proving that the fit quality increased significantly after using a bi-exponential

function and that the fit is nearly perfect in this case. Note that in the PC

mutant, full State I is not achieved because State I conditions correspond to

vigorous shaking of the sample in darkness.

211

Supplementary Figure S2:

As a qualitative assessment of the relative changes of the PSI and PSII

antenna size upon State transitions, we used the intensities of the emission

bands at 685 nm and 712 nm which are respectively specific of the PSII and

PSI core complexes and their connected antenna proteins1,2. Although it

cannot be directly converted into an absolute change in antenna size, this

provides a useful and commonly resorted to assessment of the amplitude of

ST. Panel A compares typical emission spectra obtained with a WT strain

placed in State I (blue) and State II (orange) conditions, after normalization to

the 685 nm peak. These changes were quantified as the increase of the 712

nm to 685 nm emission intensities upon State I to State II transition in the WT

and PC mutant (n=9) and are shown in Panel B. Notably, this ratio was

smaller in the latter than in the former case consistent with the notion that

vigorous shaking in the dark only promotes an incomplete State I.

A B

212

Supplementary Figure S3:

Characterization of the changes in fluorescence and phosphorylation pattern

tant.

Panel A shows the changes in the fluorescence yield in State I (oxic

conditions, blue) and State II (anoxic conditions, orange) conditions. As

schematized by the black and yellow horizontal bars, the cells were dark-

adapted and then illuminated for 4 seconds, after which a saturating multi-

turnover pulse, indicated by the vertical arrow, was applied to probe the FMax

level. It remained unchanged upon the shift from State I to State II. On the

contrary the fluorescence level, F0’, measured before the illumination,

increased upon the transition to State II (anoxic conditions), witnessing the

reduction of the plastoquinone pool in the dark and thus showing that the

conditions required to promote State II were achieved. This was further

supported by the immunoblot with an anti-phosphothreonine antibody of the

-9 strains, the latter being used as a reference strain unable

to perform ST. It shows that changes in the phosphorylation pattern expected

to take place upon the transition from State I to State II did indeed occur even

in the absence of PSI. The alpha subunit of the CF0F1 ATP synthase was

used as a loading control. The increase in D2/CP43 phosphorylation has been

previously reported and is stt7-9-independent3.

A B

-atpB

25 kDa

20 kDa

15 kDa

-phospho-Thr

stt7-9 psaB

STI STII STI STII

D2/CP43

LHC

213

Supplementary Figure S4:

mutant in State I (black) and State II (red) conditions. Besides the detecting

wavelength, 820 nm versus 705 nm, the experimental condition were exactly

the same as for the measure of P700 oxidation kinetics at 705 nm. The

comparison of the two kinetics shows that the rate of P700 oxidation

increased by 1.4 fold, consistent with the results obtained when probing the

oxidation state of P700 at 705 nm thus showing that the putative contribution

of fluorescence changes at 705 nm was negligible in our conditions. 705 nm,

rather than 820 nm was used routinely because of the better signal to noise

achieved at this wavelength.

214

Supplementary discussion:

Here we would like to briefly discuss the possible reasons that may account

for the conflicting results reported here and in the two recent reports that

concluded that mobile LHCs detaching from PSII are eventually aggregated

and efficiently quench excitation energy rather than being connected to PSI.

Unlu et al.4 used fluorescence lifetimes to assess the change in the

photosystems’ antenna size and concluded that this was unaffected by the

transition from State I to State II. However, this interpretation was essentially

based on the absence of changes in the total amplitude of the decay

components assigned to either PSII or LHCII whereas PSI, strictly speaking,

was not directly resolved in these analyses. Of course we cannot rule out the

possibility that phospho-LHCs self-quench in some peculiar environmental

situations that these authors and not us would have encountered

serendipitously. Alternatively, a spontaneous drift of the cells towards State II

owing to the incomplete experimental control of the redox state of the PQ pool

(as reported here for example in the case of the PC strain) cannot be

completely ruled out inasmuch as the measure of the lifetimes in a control

case such as the stt7 strain is lacking.

On the other hand, Nagy et al.5 used an approach similar to the one followed

here based on ECS measurements. We note however, that they compared

the increase of the PSII antenna upon State II to State I transition (75%), to

the increase of the PSI antenna size upon the transition in the opposite

direction (15%). Saliently, even if, as we propose here, the changes are

complementary, this comparison would not have given the same numbers

because the two states taken as the starting reference differ (to convince

oneself, one may compute the decrease in the PSII antenna size (from State I

to State II) from the increase of 75 % reported in Nagy et al. (from State II to

State I): 1- 1/1.75= 43% which is remarkably similar to the values reported

here). This correction being made, we acknowledge the fact that 43% remains

larger than 15%. This latter figure is however not so different from that

reported here for the PC mutant with a similar approach. Possibly, in the

conditions used by Nagy et al. the plastoquinone pool remained partially

215

reduced leading, when using the ECS approach, to an underestimation of the

changes in the PSI antenna size for the reasons discussed above. Last, the

scatter we observed between biological replicates points to an intrinsic

biological variability which may have escaped the attention of the authors who

performed a relatively low number of biological replicates (n=3).

216

1 Murata, N., Nishimura, M. & Takamiya, A. Fluorescence of chlorophyll

in photosynthetic systems III. Emission and action spectra of fluorescence—

Three emission bands of chlorophyll a and the energy transfer between two

pigment systems. Biochimica et Biophysica Acta (BBA) - Biophysics including

Photosynthesis 126, 234-243, doi:http://dx.doi.org/10.1016/0926-

6585(66)90059-8 (1966).

2 Murakami, A. Quantitative analysis of 77K fluorescence emission

spectra in Synechocystis sp. PCC 6714 and Chlamydomonas reinhardtii with

variable PS I/PS II stoichiometries. Photosynth. Res. 53, 141-148,

doi:10.1023/a:1005818317797 (1997).

3 Bergner, S. V. et al. STATE TRANSITION7-Dependent

Phosphorylation Is Modulated by Changing Environmental Conditions, and Its

Absence Triggers Remodeling of Photosynthetic Protein Complexes. Plant

physiology 168, 615-+, doi:10.1104/pp.15.00072 (2015).

4 Unlu, C., Drop, B., Croce, R. & van Amerongen, H. State transitions in

Chlamydomonas reinhardtii strongly modulate the functional size of

photosystem II but not of photosystem I. Proceedings of the National

Academy of Sciences of the United States of America 111, 3460-3465,

doi:10.1073/pnas.1319164111 (2014).

5 Nagy, G. et al. Chloroplast remodeling during state transitions in

Chlamydomonas reinhardtii as revealed by noninvasive techniques in vivo.

Proceedings of the National Academy of Sciences of the United States of

America 111, 5042-5047, doi:10.1073/pnas.1322494111 (2014).

217

Perspectives, analysis of discrepanicies between labs

Our results clearly substantiate the view that state transitions are a process of

antenna sharing between photosystem II and I. A question then arises is why

in the previously mentioned articles the effect of PSI antenna size was not

seen? Several explanations can be proposed.

Focusing first on the work of Nagy et al., there are 2 experiments that can be

discussed. Because the remodeling of the thylakoid membranes upon state

transitions, as evidenced by SANS and circular dichroism measurements, is

not debatable, solely the ECS measurements of PSI antenna size changes

are in disagreement with our data. Surprisingly, these experiments were made

with the same setup and in very similar biological conditions. The only

possible explication for the discrepancy is therefore the number of biological

replicates used – we have in some instances also obtained values, which are

lower than the mean. Secondly, the biochemical preparations, which for the

authors indicate little association of LHCII with PSI-LHCI supercomplexes, are

not conclusive because of the artificial environment used (i.e. detergent

presence), and cannot be extrapolated to the in vivo situation.

With regards to the articles exploring the fluorescence lifetimes, we will use

the Unlu et al. (2014) paper as an example of why it is difficult to draw

conclusion from in vivo measurements with this method. Because lifetimes’

analysis is not known to the author, a warm acknowledgement is necessary

for Stefano Santabarbara for the help with understanding the issues

surrounding the paper.

In the paper by Unlu et al. (2014) State Transitions were investigated in whole

cells of C. reinhardtii using time resolved fluorescence techniques, in

particular time correlated single photon counting. Cells were locked either to

state 1 or state 2 before the measurements and the kinetics of excited state

decay were recorded under conditions of “open” centres, i.e. close to F0, and

monitored at a few emission wavelengths (680, 701 and 720 nm) upon

preferential excitation of either Chl a (440 nm) or Chl b (465 nm). This

approach should allow to discriminate changes in the absorption cross-section

due to reconfiguration of external antenna, as the 465 nm excitation will

218

preferentially excited light harvesting complexes (LHC) with respect to core-

complexes.

The authors described the decay of STI and STII-locked cells, irrespectibly of

the excitation with 4 decay components which values depend on the

excitation and emission conditions. It was shown that in particular the fastest

lifetime of 66-70 ps attributed solely to PSI emission exhibited very limited

changes upon state transitions, both in terms of the lifetime and its amplitude.

Since the relative amplitude is proportional to the absorption cross-section,

the absence of variation suggested that changes in the absorption area of PSI

were little upon state transitions.

Instead, the authors proposed that changes in fluorescence intensity were

associated with a dark quenching process, likely associated with PSII antenna

aggregation. This proposition stemmed from the analysis of remaining lifetime

components from fitting of the fluorescence decay kinetics: The authors

noticed that under State 1 conditions components with lifetimes longer than

about 65 ps varied from 250 ps, 665 ps, and 1.9 ns (440 nm excitation) or 260

ps, 840 ps, 1.9 ns (465 nm excitation), to 215 ps, 550 ps and 1.2 ns (440 nm)

and 259 ps, 715 ps and 1.2 ns nm (465 nm) upon transition to State 2. They

have then estimated, assuming that all of - and only - lifetimes above 100 ps

describe PSII emission, the average lifetime using following formula:

/av i i i

i i

A A where iA are the amplitudes and i the lifetimes.

This calculation yields av values of 448-550 ps in State 1 and 446-510 ps in

state 2 conditions, for Chl a preferential excitation at 440 nm, and av between

653-666 ps in State 1 and 597-640 ps for State 2 for Chl b excitation at 465

nm. Note that interval depends on the emission wavelength, and that the

values reported in the paper refer to the 680 nm only. Therefore, the value

decrease of about 7% was reported and a quenching interpretation of the data

is the most straightforward.

A more thorough analysis indicates that larger variations are observed for the

slower component, characterised by 550-750 ps and 1.2-1.9 ns lifetimes. The

219

latter exhibit the biggest change of all, as its value almost halves during state

I-to-State II transition. It is however necessary to mention that its amplitude,

generally small and maximally of 8-15% of the total, increases in State II

conditions. This increase more than compensates the decrease of the lifetime,

because its contribution to the total amplitude reaches 30-40% for the 680 nm

excitation, resulting in an overall higher contribution of the long-lived

component in State II. The authors interpret these findings as an increase in

the pool of uncoupled PSII antenna in State II. This is logical with regards to

their previous assumptions – an antenna aggregation-related quenching

process decreases the lifetime, while its amplitude increases because more

antenna are uncoupled when the PQ pool is reduced in anoxia.

However, long lifetimes are often interpreted in the literature as resulting from

a fraction of photosystems being in their closed state, i.e. at FMax. This is due

to a non-negligible actinic effect of the excitation light. Hence they are

commonly excluded from the av computations in order not to skew the values

supposedly being measured for open photochemical centers close to the F0.

When this correction is taken into account, the value of av appear to

decrease upon State 2 conditions to 388-395 ps from the value of 455-480 ps

in State 1 (440 nm excitation). This effect is therefore even larger than when

the longer lifetimes are taken into account and amounts to a ~20% decrease.

The latter is in broad disagreement with the steady-state measurements. It is

known that the F0 value rises under reducing conditions due to the reduction

of PQ pool and the equilibrium of the latter with the QA quinone, yielding the

primary stable PSII acceptor reduced and as a result the photochemical trap

closed and in its FMax state. Unfortunately, since the authors did not report an

analysis of the excited state at FMax, it is impossible to extract detailed

comparisons concerning the FV/FM values in the two states investigated. It is

nonetheless to be expected that, for 10-20% (non-photochemical quenching)

at F0, a much more pronounced decrease in emission intensity would

manifest at FM, which will be in line with value previously reported for C.

reinhardtii of 30-40% decrease. Yet, the decrease in FV/FM would only be of

about 20-30%, which is instead much less than observed in several previous

220

studies. The reason for these discrepancies, i.e. the decrease rather than

increase in the steady-state F0 level and the relative limited decrease in FV/FM

(although the latter value is only an estimate) remain to be understood.

One could convince itself about a possible reason for such behavior in a

following manner: typical relative fluorescence induction curves in State I and

II treated with DCMU are presented. However, in order to satisfy the data

interpretation provided by the authors, the State I curve would need to present

kinetics different than the typical induction, because F0 value as measured

with the fluorescence decay method is lower in State II than in State I.

If prompted to speculate, we could hypothesize that due to the actinic effect of

the laser excitation the initial “F0” was in fact higher than the minimal value,

and/or that due to a protocol necessitating several minutes of illumination the

State II sample escaped the overreduction and its “F0” value was decreased

with regards to its initial state expected to be close to the FMax.

Finally, analysis of excited state decay in vivo, where PSI and PSII are

present simultaneously is notoriously cumbersome. Although the total decay

kinetics are described by 4 or 5 exponentials, their assignment is not

straightforward. This is because even for a homogenous, biochemically

isolated PSI-LHCa or PSII-LHCII supercomplexes their decay is

multiexponential. Typically, at least two, but more often 3 or 4 lifetimes, are

required to describe the fluorescence decay in PSI-LHCa. These lifetimes

follow 4-8 ps, 10-15 ps, 20-40 ps and 80-120 ps kinetics, the exact values

depending on the preparation. Similarly for PSII-LHCII at least 3 components

are frequently observed, falling in the 40-60 ps, 200-400 ps and 500-700 ps

ranges, again depending on the preparation and species from which the

photosystem was extracted. It is thus clear from this comparison that some of

the lifetimes overlap, and despite PSI emitting less fluorescence than PSII,

the possibility of resolving the two simultaneously is a difficult task.

The description obtained by Unlu and colleagues is not unprecedented, but

noticeably fails to resolve the fastest phase of PSII-LHCII relaxation almost

completely. It is therefore not unlikely that the single component, which is

assigned to PSI, that is somewhat longer than the values observed in vitro,

221

contains a fractional contribution of PSII decay. This has a direct effect on the

conclusions drawn about State Transitions mechanism - because of an

increase in the mixing between PSII and PSI contributions, due to a larger

impact of the latter, for instance associated to its increase in optical cross

section, would result in what would then be an apparent decrease in PSII

attributed lifetimes nullifying the eventual differences. Lastly, a hint that this

mixing could have indeed occurred during the attribution of the lifetimes is

provided by the fact that the av seems to exhibit a spectral dependence, a

feat virtually non-existent in biochemical preps of isolated photosystems.

222

Conclusions and perspectives

During the course of my PhD, I was interested in the transter of electrons in

photosynthesis and mechanisms allowing its regulation. The number of

processes and their complexity – especially in an in vivo study – make it

challenging to experimentally distinguish one form another. Moreover, due to

the design of the physiology of electron transfer, that is a multitude negative

feedback loops and multiple mechanisms of little apparent relevance this

study proved to be notoriously difficult. We have concentrated on two of them

– chlororespiration and state transitions, only to find out that one cannot

quantitatively measure chlororespiratory flux without adjusting it to the

antenna size of PSI, influenced by state transitions, and without taking into

account cyclic electron flow, a prominient alternative transfer pathway.

Similarily, physiology ot the chloroplast with notably glycolysis taking place

inside this organelle, together with a tight connection between chloroplast and

mitochondrium and the presence of a respiratory chain inside thylakoids

themselves forced us to rely on mutants blocked in the electron transfer in

order to reliably, quantitatively assess the extent of state transitions in

Chlamydomonas. Finally, a chlororespiratory mutant proved to be useful for

the validation of our hypotheses about cyclic electron transfer.

Taken together, these observations point out the difficulties that will be

encountered during future studies about the regulation of electron transfer in

photosynthesis, and great care that must be taken with regards to the

experiment design. What are the questions we think will need to be asked in

the field in the next years?

Starting with the respiratory flow in thylakoids, it is necessary to ask why it

was retained in evolution. It is clear from phylogenetic evidence for both

PTOX and a PQ reductase (NDH or NDA2) that these enzymes are

conserved in photosynthetic lineages, yet apart from our findings, roles

proposed for either PTOX or NDH/NDA2 are incompatible with their catalytic

rates.

In higher plants, the presence of the NDH is particularly puzzling. This

multimeric enzyme, with many additional subunits with respect to its

223

mitochondrial homologue (Peltier et al, 2016) is extremely slow and is not

contributing to CEF (Joliot & Joliot, 2002; Trouillard et al, 2012b); its substrate

is proposed to be ferredoxin, suggesting that instead of a chlororespiratory

role in darkness – where Fdred is absent - it is only active in the light. Until

now, apart from genetic evidence from the Shikanai group, not a lot is known

about NDH’s involvement in the photosynthetic electron flow. Is it possible

that such a huge complex serves only a regulatory or signaling role? What is

a reason for such a cost-ineffective system to be retained?

In darkness, the PQ pool of higher plants is almost completely oxidized.

Because the rate of PTOX is very low in plants, it can be hypothesized that in

fact no chlororespiratory electron flow exists in these organisms. This is

supported by the fact that NDH is an apparent Fd:PQ and not NAD(P)H:PQ

oxidoreductase, but experimental proofs are needed in order to conclude

wheter this is indeed the case. This is furthermore compatible with a different

metabolism of carbohydrates in higher plants with regards to algae – in the

former, glycolysis is entirely cytosolic and not taking place in the chloroplast.

On the other hand, we have shown that in algae, chlororespiration plays a

crucial role in the regulation of the redox status of chloroplast stroma – but

only in conditions where the metabolism contributes to the reduction of PSI

acceptors. What is a reason for retaining such a complicated process of

NAD(P)H oxidation which is entangled with the photosynthetic flow? We now

have evidence for the flavo-diiron proteins, soluble electron transporters

present in the stroma which oxidize the PSI acceptors and could readily

replace the oxidative role of chlororespiration assuming that in agreement with

our data, neither NDA2 nor PTOX are significant in the light. Is it possible that

chlororespiration has multiple contributions to plastid physiology?

If however a role in the light is envisagable for the PTOX in adult plants and

algae, it is potentially this of signaling. Due to the mechanism of oxygen

reduction proposed for alternative oxidase, PTOX being its homologue was

thought to produce reactive oxygen species, a class of radicals potentially

involved in retrograde signaling. It was experimentally confirmed that it is

indeed the case (Feilke et al, 2014; Galzerano et al, 2014; Heyno et al, 2014;

224

Yu et al, 2014) – and even more so in PTOX was overexpressed. This puts,

again, the potential role of PTOX as an electron sink in doubt.

On the other hand, due to the way in which the regulation of excitation

balance between photosystems works, chlororespiration seems to be a robust

oxygen and metabolism sensor for the photochemistry control. In case of an

influx of reductants, both the NDA2 substrate will become more abundant,

and oxygen partial pressure will decrease (as mitorespiration will increase) –

both leading to an overreduction of the PQ pool. This in turn activated the

kinase responsible for LHCII phosphorylation and state transitions. Such

qualitative signalling, rather than quantitative oxidative process seemed to

have been overlooked before (Houille-Vernes et al, 2011). We have used the

“lock” in state two of the PTOX2 mutant as a screening proxy, however an

oxygen sensor role per se was not envisaged in the mentioned publication. It

is interesting to speculate whether the algae retained PTOX and NDA2

connecting the metabolism, an oxygen probe and photosynthesis at a cost of

them slowly consuming NAD(P)H. Cultures in fluctuating oxygen (and light?)

concentrations could potentially reveal a growth phenotype of

Chlamydomonas state transition AND chlororespiration mutants, until now

only seen in a double stt7 strain combined with a respiratory chain mutation

(Cardol et al, 2009).

Both PQ reductases, NDA2 and NDH despite their low rates were proposed to

sustain a process, which has the same maximal rate as the linear electron

flow. Although Cyclic Electron Flow has been known for over half a century

(Whatley et al, 1955), and is crucial for the photosynthesis, recent discoveries

seem underwhelming and largely lack mechanical understanding of the

process. Moreover, they vastly focus on the so-called “NDH-mediated” cyclic,

most probably of low physiological importance bar for the C4 plants (Joliot &

Johnson, 2011; Joliot & Joliot, 2002; Trouillard et al, 2012b), or on the role of

auxiliary CEF proteins (Nandha et al, 2007) such as pgr5/pgrl1. It is crucial to

establish reliable experimental conditions in order to obtain access to maximal

rates of CEF in order to elucidate the actual mechanism of this process – a

feat achieved in plants (Joliot & Johnson, 2011) but not in Chlamydomonas.

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In Chlamydomonas, cyclic electron flow is even more a mystery. Because of

the absence of the red-shifted chlorophylls in Chlamydomonas PSI, its

absorption spectrum is only very slightly red-shifted with regards to higher

plants. This, in turn, yields the use of far-red light as preferentially exciting PSI

limited, while it is widely employed for the probing of the maximal CEF rates in

plants during dark-to-light transitions. Lack of this tool for the alga forced

researchers to use a different protocol – measurements of steady-state cyclic

by ECS or P700 measurements in the presence of DCMU. The rationale

behind this approach is that DCMU blocks the linear electron flow, so only the

cyclic pathway will be responsible for the reduction of P700+. It was shown

that in normal conditions CEF in Chlamydomonas operates with a rate of 10 e-

/s/PSI (Alric et al, 2010a; Takahashi et al, 2013) in the absence of LEF.

Although it is necessary to mention that some groups start to ask questions

about the relevance and rates of CEF in physiological conditions using

simultaneous measurements of PSI and PSII electron transfer rates (Fan et

al, 2016; Godaux et al, 2015a), this approach aims more at an elucidation of

physiological role of the process more than its mechanism. What can be

suggested for the future experiments aiming at elucidation of the mechanics

of CEF in vivo?

First, we need to disagree with the approach of blocking LEF if any

quantitative conclusions about CEF are to be made in the steady state.

Blocking the PSII activity results in an oxidation of the whole electron transfer

chain due to the lack of reductants. Because of that, assuming the CBB cycle

operates at its normal rate, i.e. was activated by the illumination even in the

presence of DCMU, there is a significant lack of CEF substrate (most

probably reduced Ferredoxin) making any precise measurements of the Vmax

irrelevant. This applies to the experiments aiming at establishing the maximal

rate of CEF – the only mean allowing further elucidation of the mechanism

and actors necessary for the presence of CEF, and to all experiments

involving mutants of proteins proposed to be implied in CEF.

This view is sustained by another observation, that CEF rates are higher in

anoxic conditions (Alric et al, 2010a; Clowez et al, 2015; Takahashi et al,

2013). In this case, because of halted oxidative phosphorylation but

226

ongoing/enhanced glycolysis, the quantity of reducing equivalents in

mitochondira and chloroplasts increases (note that also chlororespiration is

blocked when deprived of oxygen). Because NAD(P)H pool is virtually

completely reduced (and it is in equilibrium with ferrodoxin, also reducing at

least a part of its pool), upon illumination of a DCMU-treated sample the pool

of Ferredoxin is also more reduced than in oxic conditions. Thanks to this, in

our interpretation, the CEF measured in these conditions is faster than

normally; although it does not exclude the hypothesis of a redox-controlled

“biological” CEF activation somehow promoting this pathway for instance by

superstucturation, it is in our view no longer necessary to evoke anything else

than an abundance of CEF substrate (and its relatively slow consumption) in

anoxia. Furthermore, it was shown that this anoxia-induced CEF rate increase

is fast, pointing toward a passive and not contrived control – however due to

methodological issues CEF was measured at least after a few minutes in

anoxia (Clowez et al, 2015). Importantly, CEF is a process necessitating two

substrates, apart from Fdred also oxidized plastoquinone. In anoxia in the

presence of DCMU, upon a few seconds of illumination PSI still exhibits

overreduction-related acceptor side limitation while it is partially oxidized,

hence one can assume that due to the difference in midpoint potentials the

PQ pool is vitually completely oxidized. This favors CEF - as both of its

substrates are present - yet despite such ‘proper’ experimental setup in vivo

the CEF rates reported for Chlamydomonas scatter from 40 to 60 e-/s/PSI

(Alric et al, 2010a; Clowez et al, 2015; Takahashi et al, 2013). As pointed in

the discussion of Clowez et al (2015), the assumption that both PQ and

NAD(P)H/Fd pools need to be oxidized and reduced, respectively, is not a

necessity for an efficient cyclic – however if one follows this logic, sustaining

the maximal possible rate of the process is necessary. Hence, it is essential to

keep the light intensity - translating the kOX of the PSI which is controlling the

rate at which the PQ are oxidized - saturating (as high as possible with

regards to the rate of limiting processes). This feat was not applied in any of

the previous anoxic studies and could in fact explain the differences between

various reports. However, the use of anoxia is cumbersome and can lead to

discrepancies due to differences in the metabolic state of cells, the latter

being difficult to control for in vivo measurements.

227

Finally, differences between rates of CEF in higher plants and

Chlamydomonas raise questions about the nature of this process. Is it

possible that they are indeed so different and that the Vmax of CEF varies 2 to

3 times between algae and plants? Furthermore, despite many mutant

screens in photosynthetic eukaryotes not a single one was found to have

normal linear electron flow but no cyclic, pointing to the idea that the number

of actors involved in CEF is very limited with regards to LEF – if this is indeed

the case, then why should the cyclic photophosphorylation be different

between different lineages as the LEF actors are essentialy the same?

We focused a part of the PhD to develop a reliable, quantitative method,

which could potentially help in obtaining CEF mutants – they are convenient

to study in Chlamydomonas because if they happen to be non-

photoautotrphic they can still be easily manipulated in heterotrophic media.

The proof of concept was provided in the manuscript for the FtsH

protease/chaperon mutant, which shows extremely diminished CEF with our

method, but not with the ususal methods (A. Malnoe, PhD thesis). Conversly,

it would be telling if with this approach no new CEF-but-not-LEF mutants were

to be be found, further implying that it is the Qo site acting as the Fd-PQ

oxidoreductase (FQR) and “minimal” CEF actors are common to both CEF

and LEF.

If one thinks about CEF as a simple re-wiring device, which returns electrons

back to the PQ pool, how can it be regulated? In other words, how to control

the partition of electrons from the node – ferredoxin - going through the CEF

and LEF pathways? The obvious idea, especially supposing that CEF adjusts

the ATP/NADPH ratio, would be a positive-negative feedback: ATP enhancing

Fdred oxidation via the CBB cycle, NADPH increasing CEF due to a decrease

in capacity of Fdred oxidative pathway; or inversely, ADP increasing CEF, and

NADP+ LEF capacity.

Such passive control, however, can probably be ruled out, at least in our

experimental conditions leading to an increased CEF in fluctuating light. In the

PTOX2 mutant, as we interpret it, the overreduction of stroma in darkness

results in an enhanced CEF rate in the light and at the end of the 30 min

228

treatment. It is easy to hypothesize that the two are directly related, yet a

careful examination of the PSI acceptor limitation at the beginning and at the

end of the light period reveals that in the mutant both the quantity of

photooxidable P700 and quantity of oxidized P700 accumulated in the light

are much bigger than in the WT. This being the case, one needs to add a

long-lived regulation of CEF to the picture – a regulation of an unknown origin.

This regulation in terms of the rewiring would increase the probability that an

electron comes back to the membrane despite having a positive “hole” in the

linear pathway. How can one achieve that? If the transfer through CEF or LEF

pathways – for instance, to a b6f- of PSI-bound FNR, respectively – from

ferredoxin is not diffusion-limited, an increase in the rate of forward electron

transfer (or decrease of the rate of back-reaction) is necessary to explain any

apparent increase in CEF. Hypothetically, this could be achieved by any post-

translational modification or even a conformation change of any of the

electron carriers or other proteins present in proximity of CEF cofactors. On

the other hand, if Fdred needs to diffusionally explore a volume and then

donates the electron with a given probability to CEF or LEF pathway, another

possibility can be taken into account. This would be a structuration allowing

an increase in the frequency of Fd encountering CEF pathway with regards to

LEF – or, inversely, moving away of LEF components from PSI acceptor side.

Such structuration was apparently observed in Chlamydomonas in anoxia

(Iwai et al, 2010a) when b6f complexes were found to be physically associated

with PSI-LHCI supercomplex. In higher plants, it was shown that cyt. b6f can

be copurified with FNR, in which case the latter could serve as a bait for Fdred

in order to facilitate or increase the probability of electron donation to the

stromal haems of the b6f (Joliot & Johnson, 2011). However, the

supercomplexes’ formation, in order to be controllable, also needs some

degree of regulation, such as reversible phosphorylation or similar of at least

one of the components. We have already started to explore the biochemical

nature of the increased CEF in the PTOX mutant in fluctuating light to

elucidate whether a b6f-PSI supercomplex hypothesis is plausible, or whether

similar to higher plants it could be the FNR that modulates the repartitioning of

electrons from the Fd node.

229

In the latter case, there is a need of experimental proof that Fdred can

efficiently reduce ci or bh hemes – and possibly, for the tunneling distance to

allow a rapid transfer (Moser et al, 1992), a conformational change could be

necessary in the b6f complex (D. Picot, personal communication).

In Chlamydomonas, several different isoforms of ferredoxin exist. They are

thought to have slightly different redox potentials and were thus proposed to

have specific functions, i.e. some would serve as CEF and some as LEF

substrates (L. Mosebach, M. Hippler, personal communication). However,

ferredoxin 1 is by far the most abundant of them, and its transcript constitutes

99% of all of the Fd transcripts. Moreover, even if such specialization took

place, an eventual regulatory role for CEF is difficult to reconcile with the slow

rate of gene expression taking place in tens-of-minutes timescale.

Finally, questions about the homogeneity/heterogeneity of CEF can be asked.

It is plausible that not all of the PSI/ b6f couples participate in CEF and LEF to

an identical extent, and that part of them mediate mostly LEF, and another

part CEF. Although this adds to the complexity of the already challenging CEF

measurements, it is crucial in order to reveal regulatory mechanisms behind

this process. Fitting the reduction of P700 or our multiple ECS kinetics during

P700 oxidation with several exponentials could potentially provide a hint to

this question.

With regards to state transitions, their result (the PSI and PSII cross-section

changes) was convincingly described in our paper. However, several

questions still exist with regards to the exact mechanism by which the

transition is triggered, and how the antennas move in the tightly packed

thylakoid membrane.

It is known that a quinol presence at the QO site of the cyt. b6f activates the

stt7 kinase (Zito et al, 1999), and that it can be copurified with the b6f.

However, the sensing mechanism of the redox state of QO is still largely

unknown and only recently it was proposed that a conformational change in

the stromal loop of cyt. b6f could be sensed by the stt7 (J. Alric, 17th

International Congress on Photosynthesis Research). It is unknown how this

protein could then phosphorylate the antenna in distant regions of the

230

membrane or whether it uses some effectors performing this task or the

phosphorylation is direct.

It is clear that there is a lot of remaining questions in the field of the regulation

of electron transfer in photosynthesis.

231

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Photosynthesis and chlororespiration - competition or synergy?

Chlororespiration was initially described in Chlamydomonas reinhardtii. This

electron transfer pathway, which is found in all photosynthetic lineages,

consists of the action of a NAD(P)H:plastoquinone oxidoreductase and a

plastoquinol oxidase (PTOX). Hence, because it uses plastoquinones for

electron transport, chlororespiration constitutes an electron pathway

potentially antagonistic to the linear photosynthetic electron flow from H2O to

CO2 However, the limited flow these enzymes can sustain suggests that their

relative contribution, at least in the light and in steady-state conditions, is

limited. I thus focused on the involvement of PTOX in Chlamydomonas during

transitions from dark to light and vice versa. I found that, following a brief

illumination, the redox relaxation of the chloroplast in the dark was much

affected when PTOX2, the major plastoquinol oxidase in Chlamydomonas, is

lacking. Importantly, I show that this has a significant physiological relevance

as the growth of a PTOX2- lacking mutant is markedly slower in intermittent

light, which can be rationalized in terms of a decreased flux sustained by

photosystem II. I also investigated the influence of chlororespiration on cyclic

electron flow using novel experimental techniques combined with theoretical

modelling. Last, I explored, in collaboration with Stefano Santabarbara, the

mechanism for redistribution of light excitation energy between the two

photosystems, a process triggered by changes in the redox state of

plastoquinone pool. I thus showed that, contrarily to what has been suggested

recently, this regulation mechanism corresponds to an actual transfer of light

harvesting antenna between the two photosystems.

Keywords: photosynthesis, respiration, state transitions, electron transfer,

regulation, cyclic electron flow

Photosynthèse et chlororespiration - compétition ou synergie?

La chlororespiration a été initialement décrite chez Chlamydomonas

reinhardtii. Cette voie alternative du transfert d’électrons, présente dans

toutes des lignées photosynthétiques, est constituée par l’activité d’une

NAD(P)H:plastoquinone oxidoreductase, et d’une plastoquinol oxydase

(PTOX). Parce qu’elle utilise les plastoquinones comme transporteur

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d’électrons, la chlororespiration représente une voie potentiellement

antagoniste au transfert photosynthétique linéaire de l’eau au CO2.

Néanmoins, le faible flux autorisé par ces enzymes suggère que, au moins

sous éclairement continu et en conditions stationnaires, leur contribution est

limité. Je me suis donc concentré sur la rôle du PTOX pendant les transitions

lumière-obscurité et vice-versa. J’ai observé qu’après une brève illumination,

la relaxation redox du chloroplaste est entravée quand PTOX2, l’oxydase

chlororespiratoire majeure chez Chlamydomonas, est absente. J’ai démontré

la pertinence physiologique de cette observation par une étude comparative

des courbes de croissance de souches mutantes pour PTOX et de la souche

sauvage :la croissance du mutant de PTOX2 est dramatiquement retardée en

condition de la lumière intermittente – ce qui peux être expliqué par une

diminution du flux d’électrons à partir du photosystème II. Je me suis

également intéressé à l’impact de la chlororespiration sur le flux d’électrons

cyclique en utilisant une nouvelle approche spectroscopique combinée à de la

modélisation. Enfin, j’ai exploré, en collaboration avec Stefano Santabarbara,

le mécanisme de redistribution de l’énergie lumineuse entre les deux

photosystèmes, mécanisme gouverné par des changements d’état redox des

plastoquinones. J’ai ainsi démontré que, contrairement à de récentes

suggestions, un véritable transfert d’antennes collectrices d’énergie, se

produit entre les deux photosystèmes.

Mots clés : photosynthèse, respiration, transitions d’état, transfert d’électrons,

régulation, flux d’électrons cyclique