Structure of a PSI–LHCI–cyt b6f supercomplex inChlamydomonas reinhardtii promoting cyclicelectron flow under anaerobic conditionsJanina Steinbecka,b, Ian L. Rossb, Rosalba Rothnagelb, Philipp Gäbeleina, Stefan Schulzea,1, Nichole Gilesc, Rubbiya Alib,2,Rohan Drysdaleb, Emma Siereckic, Yann Gambinc, Henning Stahlbergd, Yuichiro Takahashie, Michael Hipplera,3,and Ben Hankamerb,3

aInstitute of Plant Biology and Biotechnology, University of Münster, 48143 Münster, Germany; bInstitute for Molecular Bioscience, University ofQueensland, St. Lucia, QLD 4072, Australia; cEuropean Molecular Biology Laboratory Single Molecule Science, Lowy Cancer Research Centre, University ofNew South Wales, Sydney, NSW 2052, Australia; dCenter for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, CH-4058 Basel,Switzerland; and eResearch Institute for Interdisciplinary Science, Okayama University, 700-8530 Okayama, Japan

Edited by Krishna K. Niyogi, Howard Hughes Medical Institute and University of California, Berkeley, CA, and approved August 23, 2018 (received for reviewJune 13, 2018)

Photosynthetic linear electron flow (LEF) produces ATP andNADPH, while cyclic electron flow (CEF) exclusively drives photophos-phorylation to supply extra ATP. The fine-tuning of linear and cyclicelectron transport levels allows photosynthetic organisms to balancelight energy absorption with cellular energy requirements underconstantly changing light conditions. As LEF and CEF share manyelectron transfer components, a key question is how the same indi-vidual structural units contribute to these two different functionalmodes. Here, we report the structural identification of a photosystem I(PSI)–light harvesting complex I (LHCI)–cytochrome (cyt) b6f supercom-plex isolated from the unicellular alga Chlamydomonas reinhardtii un-der anaerobic conditions, which induces CEF. This provides strongevidence for the model that enhanced CEF is induced by the formationof CEF supercomplexes, when stromal electron carriers are reduced, togenerate additional ATP. The additional identification of PSI–LHCI–LHCII complexes is consistent with recent findings that both CEF en-hancement and state transitions are triggered by similar conditions,but can occur independently from each other. Single molecule fluores-cence correlation spectroscopy indicates a physical association be-tween cyt b6f and fluorescent chlorophyll containing PSI–LHCI super-complexes. Single particle analysis identified top-view projections ofthe corresponding PSI–LHCI–cyt b6f supercomplex. Based on molecularmodeling and mass spectrometry analyses, we propose a model inwhich dissociation of LHCA2 and LHCA9 from PSI supports the forma-tion of this CEF supercomplex. This is supported by the finding thata Δlhca2 knockout mutant has constitutively enhanced CEF.

cyclic electron flow | supercomplex | photosystem I | cytochrome b6f |Chlamydomonas reinhardtii

Photosynthesis captures solar energy and stores it in the formof chemical energy, which is essential to support life on

Earth. Photosynthetic electron transport operates in two modes:linear (LEF) and cyclic electron flow (CEF). LEF yields ATPand NADPH, while CEF exclusively drives ATP production (1).Fine-tuning LEF and CEF maintains the ATP/NADPH equi-librium and efficient carbon assimilation (2, 3). CEF also playsan important role in photoprotection (4, 5) as it maintains thenecessary ΔpH across the thylakoid membrane to allow energy-dependent nonphotochemical quenching and to control the ratelimiting step of LEF (6). The dynamic tuning between LEF andCEF is therefore essential for efficient photosynthesis.LEF involves in-series activity of photosystem II (PSII), cyto-

chrome (cyt) b6f, and photosystem I (PSI), while CEF involvesonly PSI and cyt b6f. During CEF, electrons released by PSI arereinjected into the photosynthetic electron transport chain at theplastoquinone (PQ) pool or at the stromal side of the cyt b6fcomplex. The fact that LEF and CEF share many electrontransfer components (e.g., PSI and cyt b6f) raises the question of

how these membrane protein complexes can contribute to bothfunctional modes. Extensive biochemical and biophysical analy-ses using the green alga Chlamydomonas reinhardtii suggest thatefficient CEF depends on the formation of a CEF supercomplexconsisting of PSI, cyt b6f, and subunits ferredoxin–NADP–oxidore-ductase (FNR), Proton Gradient Regulation-Like 1 (PGRL1), An-aerobic Response 1 (ANR1), and Calcium Sensor (CAS). The CEFsupercomplex is proposed to enhance CEF over LEF when stromalelectron carriers are reduced (excess NADPH) and ATP is limiting(7–10). However, structural evidence for this supercomplex in C.reinhardtii is lacking, probably due to its putative dynamic nature.Here, under CEF-inducing anaerobic conditions, a sucrose

density gradient (SDG) fraction with CEF activity (7, 8) wasisolated from C. reinhardtii, and a PSI–light harvesting complex I(LHCI)–cyt b6f-containing CEF supercomplex within it, wasstructurally characterized. The physical association betweenPSI–LHCI and cyt b6f was supported using single molecule

Significance

To optimize photosynthetic performance and minimize pho-tooxidative damage, photosynthetic organisms evolved to ef-ficiently balance light energy absorption and electron transportwith cellular energy requirements under constantly changinglight conditions. The regulation of linear electron flow (LEF)and cyclic electron flow (CEF) contributes to this fine-tuning.Here we present a model of the formation and structural molec-ular organization of a CEF-performing photosystem I (PSI)–lightharvesting complex I (LHCI)–cytochrome (cyt) b6f supercomplexfrom the green alga Chlamydomonas reinhardtii. Such a struc-tural arrangement could modulate the distinct operation of LEFand CEF to optimize light energy utilization, despite the sameindividual structural units contributing to these two differentfunctional modes.

Author contributions: J.S., I.L.R., M.H., and B.H. designed research; J.S., I.L.R., P.G., andN.G. performed research; H.S. and Y.T. contributed new reagents/analytic tools; J.S., I.L.R.,R.R., P.G., S.S., R.A., R.D., E.S., Y.G., M.H., and B.H. analyzed data; and J.S., I.L.R., M.H., andB.H. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.1Present address: Department of Biology, University of Pennsylvania, Philadelphia,PA 19104.

2Present address: Centre for Microscopy and Microanalysis, University of Queensland,St. Lucia, QLD 4072, Australia.

3To whom correspondence may be addressed. Email: mhippler@uni-muenster.de orb.hankamer@uq.edu.au.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1809973115/-/DCSupplemental.

Published online September 25, 2018.

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fluorescence (SMF) correlation spectroscopy. Immunoblot andmass spectrometry (MS) analyses also clearly identified PSI,LHCI, cyt b6f as well as FNR, PGRL1, ANR1, and CAS. Theirstructural organization was characterized using crosslinking, MS,and single particle analysis (SPA). In Chlamydomonas PSI–LHCI,the LHCA2 and LHCA9 subunits are located at its PSAG/H side(11) similar to the recently resolved PSI structure of a red alga (12).Our CEF data suggest a dynamic dissociation/association model, inwhich LHCA2 and LHCA9 dissociate from PSI–LHCI enablingCEF supercomplex formation.

ResultsIdentification of a PSI–LHCI–cyt b6f Supercomplex. CEF super-complexes of C. reinhardtii were isolated from anaerobically cul-tured cells (7, 8). Isolated thylakoid membranes were solubilizedwith n-dodecyl α-D-maltoside (α-DDM) and fractionated usingSDG centrifugation (7, 8). Immunoblot analysis identified a highmolecular weight SDG fraction containing the major CEF super-complex components PSI and cyt b6f (Fig. 1 B and C). Previouswork had demonstrated CEF activities in the same SDG fractions,but it remained possible that this was due to (i) colocalization of cytb6f and PSI in small residual membrane patches (since α-DDM is amild detergent) or (ii) comigration of separate cyt b6f and PSIsupercomplexes on the SDG (e.g., due to the presence of othermolecular partners such as LHCII trimers, ATPase dimers, orNDH). We therefore used SMF correlation spectroscopy (SI Ap-pendix, Figs. S1–S3) to demonstrate a single molecule, physical in-teraction between cyt b6f and PSI–LHCI complexes (Fig. 2 and SIAppendix, Fig. S3; see SI Appendix for more details). Total fluo-rescence profiles across the SDG fractions showed that the loca-tions of cyt f (labeled with green DyLight 488), as well as PSI

and PSII in the SDG correspond well with increases in greenfluorescence (Fig. 1 A and C) and red chlorophyll fluores-cence signals (Fig. 1 A and B), respectively. The fraction 19–22 pool was repurified (Fig. 1 and SI Appendix, SI Materialsand Methods) via a subsequent SDG (Fig. 2B) and measuredby SMF correlation spectroscopy to discriminate betweensingle (red or green) vs. coincident (red and green) fluores-cent events (SI Appendix, Fig. S2 B and C) corresponding todiscrete PSI–LHCI and cyt b6f molecules vs. associated PSI–LHCI–cyt b6f supercomplexes (13, 14). The frequency ofSMF coincident events (Fig. 2A) in the numbered fractions ofthe repurified complex gradient (Fig. 2B) clearly revealed thatDyLight-tagged cyt b6f is physically associated with a chloro-phyll fluorescent complex in subfractions 23–27, consistentwith the location of cyt f and PsaC (Fig. 2C). Coincidentevents were substantially higher than in the control fractionsmeasured and exceeded the number of coincident peaks ofvery high molecular weight fractions (e.g., fraction 14), inwhich broad SMF peaks were occasionally visible due to thepresence of small membrane patches (SI Appendix, Fig. S3B).Importantly, these membrane patches contributing to falsepositive coincident events were not visible in fractions 23–27,reinforcing the conclusion that the observed coincidence isdue to genuine CEF supercomplexes. Although an interactionof cyt f with PSII cannot be excluded based on these mea-surements alone, functional interaction between PSI–LHCIand cyt b6f has been reported (7, 8, 10), while a PSII–cyt b6fsupercomplex is unprecedented. Furthermore, immunoblot-ting and electron microscopy (EM) confirmed that the con-centration of PSI in these fractions was approximately 10times that of PSII (see below). Collectively these resultsprovide evidence for a physical association between PSI–LHCI and cyt b6f essential for the formation of CEF super-complexes in solution.

Fig. 1. Identification of the CEF supercomplex peak fraction in SDG by SMFspectroscopy and Western blot analysis. (A) Fluorescence intensity screeningof each SDG fraction (log scale) confirms good correlation of red chlorophyllfluorescence with the location of the photosynthetic complexes in the SDG(B) and reveals two main peaks of green fluorescent DyLight 488–trisNTAlabeled cyt f. As control for the green fluorescent signal, a SDG of an un-labeled cyt f His-Tag strain was screened (displayed in gray). (B) SDG of an-aerobic α-DDM solubilized cyt f His-Tag thylakoids separated into 77fractions. (C) Immunoblot detection of cyt f, the PSI subunit PSAC, and thePSII subunit PsbA D1 show that the PSAC signal peaks with the higher mo-lecular weight green fluorescent peak signal of the DyLight 488–trisNTAlabeled cyt f at fraction 20.

A

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Fig. 2. Cytochrome b6f is physically associated with chlorophyll fluorescentproteins in the CEF supercomplex sucrose density region revealed by SMFcoincidence analysis. (A) Coincident events, i.e., the simultaneous bursts ofgreen and red fluorescence, indicative of the physical association of DyLight488-labeled cyt f and chlorophyll fluorescent proteins, are most abundant inthe CEF supercomplex region of the SDG. The frequency of coincident eventsrelative to total fluorescent events recorded over a period of 60 s is plottedfor selected fractions over the corresponding SDG. A false positive rate of5% was applied to exclude the possible random excitation of two singlefluorescent proteins as experimentally examined previously (13). (B) PooledCEF supercomplex fractions from five SDGs of a cyt f His6-tag strain wereconcentrated, labeled with DyLight 488–trisNTA, and ultracentrifuged on asubsequent SDG to enrich for potential CEF supercomplexes. (C) Immunoblotdetection of cyt f, PSAC, and Psba D1 over the SDG fractions confirms thatthe highest frequency of coincident events correlates with the localization ofthese proteins in the high molecular weight CEF supercomplex region.

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Structural Characterization of the CEF Supercomplex. Initial singleparticle analysis revealed heterogeneity within the PSI–LHCI–cyt b6f supercomplex SDG fraction (SI Appendix, Fig. S4).Consequently, subsequent EM preparations employed an addi-tional gentle affinity purification step using a His-tagged PSI–LHCI (PSAA His-Tag) construct (15). To stabilize this super-complex, chemical protein crosslinking was performed duringthe solubilization step before affinity purification (Materials andMethods). This enabled the identification of putative CEF super-complexes by transmission electron microscopy (TEM) SPA. Atotal number of 2,708 micrographs of negatively stained particleswere recorded and yielded a total of 526,519 projection images ofprotein complexes (SI Appendix, Fig. S4B). Using SPA with Relion(16), these particles were classified into 350 classes (SI Appendix,Fig. S4C). Parallel liquid chromatography (LC)-MS/MS analysiswas conducted (SI Appendix, Fig. S8) and it identified contaminantsincluding mitochondrial NADH dehydrogenase (79,709 particles),mitochondrial ATPase (26,704 particles), and PSII–LHCII com-plexes (14,664 particles), and the corresponding SPA classes wereidentified (SI Appendix, Fig. S5) and removed from the dataset,yielding 160,819 potential PSI–LHCI-containing particles (SI Ap-pendix, Fig. S6A). A subset of 52,316 particles were identified astop-view projections of single PSI–LHCI complexes (50 classes, seeSI Appendix, Fig. S6B), as opposed to 12,179 top-view projectionsthat contained additional density adjacent to the PSI–LHCI com-plex (SI Appendix, Fig. S6C), potentially consisting of cyt b6f orLHCs. Next, tilt views and PSI–LHCI complexes lacking well-connected additional densities were eliminated from this dataset,yielding 1,139 particles. These were classified into 16 classes (SIAppendix, Fig. S7) and analyzed (Fig. 3) to confirm the ability todistinguish between PSI–LHCI–cyt b6f supercomplexes (Fig. 4) andPSI–LHCI–LHCII supercomplexes (Fig. 3 A–F, see Discussion).

A projection map of a PSI–LHCI-containing supercomplexrepresenting 50% of classes of SI Appendix, Fig. S7 (SI Appen-dix), is depicted in Fig. 4A. The molecular model overlay of PSI–LHCI (Fig. 4B, yellow and green, respectively) clearly identifiesthe PSI–LHCI supercomplex (17, 18) and an additional densityadjacent to its PSAG–LHCA1 side that can accommodate a cytb6f dimer (Fig. 4B, purple, ref. 19; see also SI Appendix, Fig. S9).Corresponding PSI–LHCI supercomplex controls are providedin Fig. 4 C and D. Based on this, the first top view of the dynamicCEF supercomplex of the green alga C. reinhardtii containingPSI, LHCI, and cyt b6f is proposed (Fig. 4B).Fig. 5 provides more detailed structural insights into factors

proposed to control the formation of this CEF supercomplex. ThePSI dataset (SI Appendix, Fig. S6B) included a larger PSI–LHCIsupercomplex (Fig. 5A) than is shown in Fig. 4C (see Fig. 5B, blueshading) highlighting the dynamic nature of these supercomplexes.The additional (Fig. 5B, unshaded) density on the PSAG side isattributed to LHCA2, LHCA9, and PSAH (Fig. 5C) based on refs.11 and 18. These subunits appear to have dissociated in manyisolated PSI–LHCI particles (Fig. 4 C and D) and are also absentin the CEF supercomplex (Fig. 4B). This suggests that PSAG andLHCA1 form an interface with the cyt b6f dimer (Fig. 5 C and D).

A Proposed Role for LHCA2 and LHCA9 in CEF Supercomplex Formation.The PSI–LHCI supercomplex projection map (Fig. 5A) fits wellwith the newly proposed model of Chlamydomonas PSI–LHCI(11). According to crosslinking and interaction studies, Ozawaet al. (11) concluded that LHCA2 and LHCA9 are not included inthe two LHCI layers (Fig. 5C, Bottom, green), but are located atthe other side of the PSI core (Fig. 5C, Top, green) and associatewith PSAB and PSAH. This subunit assignment fits the additionalPSI density identified here (Fig. 5 B and C). This model therefore

Fig. 3. Structural characterization of potential PSI–LHCI–LHCII supercomplexes from C. reinhardtii by single particle electron microscopy. A–C show three ofthe projections from SI Appendix, Fig. S6, which have densities additional to those of the PSI–LHCI supercomplex. (D–F) These projection maps have beenoverlaid with the densities of the PSI core complex [yellow (17), Protein Data Bank (PDB) 4Y28] and the LHCA proteins (green, PDB 4Y28). A LHCII trimer[magenta or cyan (42), PDB 1RWT] can be seen to fit well into these three densities. (G) Modeling a cyt b6f monomer [purple (19), PDB 1Q90] into the ad-ditional density next to PSI–LHCI supercomplex (A) shows a poor fit. (H) Overlay of the large PSI supercomplex projection map from Fig. 4A and a mesheddensity map of A. (I) Poor fit of two LHCII trimers (42) into the projection map from Fig. 4A next to the PSI–LHCI complex. This suggests that the additionaldensities in A–C are likely LHCII trimers, while the new density in H and Fig. 4A best fits a cyt b6f dimer. (Scale bar: 5 nm.)

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differs from the one presented in ref. 18. Importantly, quantitativeMS analysis (SI Appendix, Fig. S8) confirmed that LHCA2 andLHCA9 together with PSAH are most easily lost during purifi-cation (SI Appendix, Fig. S8A), consistent with Fig. 5. Strikingly, aΔlhca2 knockout mutant showed enhanced cyclic electron flowcompared with wild-type (WT) levels under anaerobic conditions;even under aerobic conditions, CEF rates in Δlhca2 were alreadyas high as for WT in anaerobic conditions (Fig. 5E). This suggeststhat the dissociation of LHCA2 and LHCA9 from the PSI–LHCIsupercomplex is important for the assembly of the CEF super-complex and the promotion of CEF (Fig. 5F).

DiscussionStructure of the Proposed CEF Supercomplex. In 2010, Iwai et al. (7)presented biochemical evidence for the existence of a CEFsupercomplex in the green alga Chlamydomonas. Here, thisbiochemical evidence is explained in structural terms. First, SMFspectroscopy supported the existence of discrete detergent sol-ubilized supercomplexes containing both cyt b6f and PSI–LHCIin solution (Figs. 1 and 2 and SI Appendix, Fig. S2); using MS, theCEF supercomplex preparation was shown to contain FNR,PGRL1, ANR1, and CAS (SI Appendix, Fig. S8) which are im-portant for CEF (8, 9). Second, having identified the presence ofintact PSI–LHCI–cyt b6f supercomplexes (SI Appendix, Fig. S2),over 500,000 TEM projection images of molecules in this prep-aration were analyzed by SPA (SI Appendix, Figs. S4–S7). Thisanalysis yielded a top-view projection map of the proposed CEFsupercomplex that is able to accommodate the PSI core (Fig. 4B,yellow), its eight LHCI antenna proteins (Fig. 4B, green), and a

cyt b6f dimer (Fig. 4B, purple). Third, these TEM data combinedwith functional data from LHCA2 knockouts yielded a model,which incorporates all of the above information and implies arole for LHCA2 and LHCA9 in the CEF supercomplex assemblyprocess (Fig. 5 E and F).The assembly of the CEF supercomplex when stromal electron

carriers are reduced to produce extra ATP (20, 21) places the PSI–LHCI supercomplex and the cyt b6f dimer in close proximity tofavor CEF over LEF. As electron transport via plastocyanin andferredoxin is thought to be diffusion limited (22), the regulation ofthe distances by structural alignment of the PSI–LHCI super-complex with cyt b6f could control electron transfer kinetics, asseen via regulation of thylakoid stacking in vascular plants (23).

CEF Supercomplex Assembly and the Promotion of Cyclic ElectronFlow. A surprising finding from the structural data (Fig. 5) wasthe absence of densities corresponding to LHCA2, LHCA9, andPSAH in the CEF supercomplex (Fig. 5 D vs. B). This led us toexamine the functional consequence of LHCA2 knockout onCEF. Strikingly, the Δlhca2 knockout mutant (11) showed con-stitutively high CEF rates under aerobic conditions, that exceededWT rates in anaerobic conditions (Fig. 5E). This suggestspotential roles for LHCA2 and LHCA9, whose association tothe PSI core is unstable in the absence of LHCA2 (11), duringthe CEF supercomplex assembly process: LHCA2 and LHCA9could therefore block the CEF supercomplex formation, andtheir removal could expose a binding site for cyt b6f required todrive CEF supercomplex assembly. This suggests a role for LHCsabove and beyond their light harvesting and energy dissipationfunctions. Notably, LHCA6 is important for NADP(H) de-hydrogenase–PSI supercomplex formation in Arabidopsis (24, 25),independently showing the role of a LHCA polypeptide insupercomplex formation. Closer analysis of a series of identifiedPSI–LHCI supercomplexes with additional putative bound LHCIItrimers shows that other LHC complexes could potentially com-petitively block cyt b6f dimer binding (e.g., Fig. 3H, see below).

State Transitions–CEF. Both state transitions and CEF enhance-ment are triggered by similar conditions, for example, the redoxstate of the chloroplast, although these processes can occur in-dependently of one another (8, 10). Hence, the presence ofdifferent PSI–LHCI-containing supercomplexes under anaer-obic conditions is expected. To eliminate the possibility thatthe proposed CEF supercomplex could represent a PSI–LHCI–LHCII complex, we modeled two LHCII trimers into this den-sity; these fitted poorly (Fig. 3I). While the focus of this paper ison the CEF supercomplex, it is of note that we also identifiedthree PSI–LHCI–LHCII complexes (Fig. 3) with an LHCIItrimer-like density localized at the LHCI belt side of the complex.These complexes represented the other 50% of classes in SI Ap-pendix, Fig. S7. MS data confirmed that LHCII proteins were lessdepleted by PSI His-Tag purification than PSII core subunits (SIAppendix, Fig. S8F), indicating that additional LHCIIs might in-deed be attached to PSI–LHCI. The supramolecular organizationof these PSI–LHCI–LHCII supercomplexes differs substantiallyfrom other C. reinhardtii PSI–LHCI–LHCII supercomplexes al-ready described, in which two LHCII trimers and one LHCIImonomer were attached to the PSAH PSAL side of PSI oppositethe LHCI ring (26). In contrast, in the PSI–LHCI–LHCII com-plexes identified here, an LHCII trimer localized at previouslyunreported positions at the outer LHCI belt side (Fig. 3 D–F),which are similar to a recent structure described in Arabidopsisthaliana where an LHCII trimer was associated with PSI at theside of LHCA2 and LHCA3 (27).The density assigned to the cyt b6f dimer at the PSAG–

LHCA1 side of PSI–LHCI in the newly identified CEF super-complex (Fig. 5) is located similarly to that observed in TEMimages of detergent solubilized PSI–LHCI–cyt b6f complexesfrom A. thaliana (27). In the Chlamydomonas CEF super-complex, the cyt b6f dimer interacts via its long side (Figs. 4B and5D); in contrast in A. thaliana, the cyt b6f dimer is reported to

Fig. 4. Structural characterization of a PSI–LHCI–cyt b6f supercomplex fromC. reinhardtii by single particle transmission electron microscopy. (A) Aver-aged TEM projection map of a large supercomplex consisting of PSI and LHCIwith an additional particle at its PSAG LHCA1 side (sum of 132 particles,representing 50% of classes in SI Appendix, Fig. S7). (B) Structural assign-ment of this supercomplex based on fitting with the crystal structures of thePSI–LHCI complex (17) and the cyt b6f complex (19) [Protein Data Bank (PDB)accession nos. 4Y28 and 1Q90, respectively]. The PSI core complex is shown inyellow with LHCA proteins highlighted in green. The cyt b6f complex isshown in purple. (C) Averaged projection map of a PSI–LHCI complex missingPSAH at its core. (D) Structural assignment of the PSI–LHCI complex similar toB. Eight LHCA proteins were modeled into the double-layered LHCI beltaccording to ref. 11. Arrows indicate the position of PSAG (yellow) and twoLHCA1 subunits (green). (Scale bar: 5 nm.)

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interact with PSI–LHCI via its short side (27). Since Chlamy-domonas PSI is larger than plant PSI due to higher numbers ofLHCI subunits forming a second LHCI belt and the potentiallyalgae-specific positions of LHCA2 and LHCA9 with respect tothe PSI core, differences may exist in regulation of CEF assemblyformation in plants compared with Chlamydomonas. The bindingof the cyt b6f dimer to the PSI–LHCI supercomplex positions theplastocyanin binding sites of cyt f (28, 29) and PSAF (30) close toone another, making the distance for electron transfer of plas-tocyanin between cyt b6f and PSI–LHCI at the luminal side rela-tively short. However, electron transfer partitioning between LEFand CEF is most likely regulated at the level of PQ reduction (31)and therefore located toward the stromal side. Cyt b6f dimer asso-ciated with its long side to PSI–LHCI would bring the more cen-trally located components involved in the stromal electron transfer,such as cyt b6 and Rieske subunits as well as the PSAA/PSAB re-action center and ferredoxin binding sites PSAC, PSAD, and PSAE(32, 33) into much closer proximity compared with a short side-bound cyt b6f, with cyt f and PSAF still in close proximity.Building on these advances, the next challenge is to obtain a

physically stable and pure CEF supercomplex for future atomicresolution structure determination to identify the position ofadditional small proteins in the CEF supercomplex like FNR,PGRL1, ANR1, and CAS (7–9). These subunits were identifiedin the CEF supercomplex SDG fraction (SI Appendix, Fig. S8C)and are expected to be attached to large supercomplexes (7–9) toenable them to migrate to this high molecular density. We at-tribute the low proportion of intact complexes on negative stainTEM grids to the labile nature of the supercomplex. A betterunderstanding of the role of LHCA2 and LHCA9 in CEF assemblycould lead to the production of physically stable supercomplexes,which would greatly assist in solving its atomic structure.

Materials and MethodsStrains and Culture Conditions. The C. reinhardtii strains cyt f–His-Tag (19, 34),PSAA–His-Tag (15), a Δlhca2 (35), and CC-4533 (WT) were used in this study.Cells were grown in tris acetate phosphate (TAP) media (36) (22 °C, 50 μMphotons m−2·s−1, 120 rpm shaking). For thylakoid isolation, cells were grown

to 3–4 × 106 cells mL−1 and harvested by centrifugation (4,600 × g, 5 min,25 °C), resuspending to 2 × 108 cells mL−1 in H1 buffer (25 mM Hepes-KOH pH7.5, 5 mM MgCl2, 0.3 M sucrose). Anaerobic conditions were induced using 2mg·mL−1 glucose oxidase (Aspergillus niger) and 50 units·mL−1 catalase (bo-vine liver, Sigma-Aldrich) with 100 mM glucose in the dark for 60 min (10).

Thylakoid and Photosynthetic Complex Isolation and DyLight 488–Tris–NTA Labelingof Photosynthetic Complexes. Thylakoids and photosynthetic complexes wereisolated according to refs. 37 and 38; see also SI Appendix, SI Materials andMethods. Cyt f–His-Tag was labeled with DyLight 488– Tris–NTA before solubili-zation and SDG-based purification (SI Appendix, SI Materials and Methods).

Single Molecule Fluorescence Measurements. Single molecule spectroscopy (SIAppendix, Fig. S2) was performed based on refs. 13 and 14 and SI Appendix, SIMaterials and Methods. Two excitation lasers (488 nm and 561 nm) focused insolution using a 40×/1.2 N.A. water immersion objective (Zeiss) simultaneouslyexcite green (here: the DyLight 488-labeled cyt f protein, see SI Appendix, SIMaterials and Methods for more details) and red (here: chlorophyll-containingproteins) fluorophores. Fluorescence was collected and separated using a 565-nmdichroic mirror; signal from DyLight 488-labeled cyt f protein was passedthrough a 525/20-nm band pass filter, while chlorophyll fluorescence was fil-tered by a 580-nm long pass filter. The fluorescence of the two channels wasrecorded simultaneously in 1-ms time bins for 60 s. For single-molecule co-incidence detection, the coincidence ratio was calculated as in ref. 13.

Immunoblot Analysis. A fixed volume (20 μL) of representative fractions acrossthe SDG gradient was analyzed by 4–12% Bis-Tris SDS/PAGE (Invitrogen).Proteins were transferred to PVDF Millipore membrane (Merck) with theXCell Blot II Module (Invitrogen) and blocked with skimmed milk. Themembrane was incubated with antibodies against PSAC (1:1,000; Agrisera),Psba D1 (1:10,000; Agrisera), and cyt f (1:5,000; Agrisera), and anti-rabbit IgG(HRP, 1:2,500; Sigma Aldrich) as the secondary antibody. Signal detectionwas performed with ECL (Amersham GE Healthcare).

Chemical Crosslinking. To crosslink proteins with disuccinimidyl suberate (DSS)(Thermo Fisher), PSAA His-Tag thylakoids were resuspended in Hepes buffer(50 M Hepes-KOH pH 8, 5 mM MgCl2) and freshly dissolved DSS (in DMSO,20 mg/mL stock concentration) was added to the sample (final concentration,0.15 mg/mL) before solubilization of proteins. The crosslinking reaction was

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Fig. 5. Dissociation of LHCA2 and LHCA9 from thePSI–LHCI complex favors the association of PSI–LHCI–cyt b6f supercomplexes and enhances cyclic electronflow. (A) Averaged TEM projection map of a PSI–LHCIcomplex with an additional density at its core. (B)Overlay of the PSI–LHCI complex from A with Fig. 4C.(C) Structural assignment of the PSI–LHCI complexfrom A based on fitting with the crystal structures ofthe PSI–LHCI complex (17). The additional densitiescompared with the smaller PSI–LHCI complex (B)were modeled with PSAH (blue) and two additionalLHCA proteins (green) at the PSI core according toref. 11. (D) Overlay of the CEF supercomplex pro-jection map from Fig. 4A with the PSI–LHCI complexfrom Fig. 4C. (Scale bar: 5 nm.) (E) Cyclic electrontransfer rates of a Δlhca2 mutant compared withwild-type levels in aerobic and anaerobic conditions.Rates were measured in steady state upon a transi-tion from darkness to light with ∼130 μE m−2·s−1 lightintensity. To exclude contribution of PSII to theelectron transfer, cultures were treated with 40 μMDCMU. Anaerobic conditions were achieved by ad-dition of 100 mM glucose and 2 mg·mL−1 glucoseoxidase. To alleviate PSI acceptor side limitationupon transition to anaerobiosis, anaerobic sampleswere kept in the darkness for 40 min and continu-ously illuminated for 2 min before the rate mea-surements (n = 6 biological replicates ± SD). Statisticalanalysis: one-way ANOVA followed by a Tukey testfor pairwise comparison of the means (***P < 0.001;*P < 0.05). (F) Structural model of CEF supercomplex formation upon dissociation of LHCA2, LHCA9, and PSAH from the PSI–LHCI complex. The PSI core isshown in yellow, LHCI proteins in bright green, cyt b6f in purple. Plastocyanin (cyt f in cyan, PSAF in orange) and ferredoxin (PSAD/E in red and cyt b6 in pink)-binding regions are indicated. All high-resolution components have been filtered to 20 Å to avoid overinterpretation.

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performed for 30 min at room temperature in the dark with occasional in-version and stopped by adding Tris buffer (final concentration, 15 mM pH 7.5,1 mM EDTA).

Purification of His-Tagged Proteins. His-tagged proteins were purified byimmobilized metal ion chromatography according to ref. 15 with the fol-lowing modifications. PSI–LHCI–cyt b6f containing fractions from six SDGswere pooled (∼4 mL) and loaded onto a 1-mL HiTrap HP column (GEHealthcare) preequilibrated with 5 mM tricine-KOH pH 8, 0.02% α-DDM,10 mM NaCl, 5 mM MgSO4, 0.5 M sucrose, 2 mM imidazole. The column waswashed with two washing buffers, with increasing the imidazole concen-trations (10 mM to 20 mM) and decreasing the sucrose concentration (0.5 Mto 0 M, 10 mL each). Elution was performed by increasing the imidazoleconcentration to 200 mM (4 elution fractions, 500 μL each).

Mass Spectrometry. For quantitative mass spectrometric analysis, the PSAAHis-Tag strain was isotopically labeled with 14N and 15N. The pooled 15Nlabeled PSI–LHCI–cyt b6f fractions (before His-Tag purification) were mixed1:1 with the His-Tag purified eluate of the 14N labeled PSI–LHCI–cyt b6fsample. Duplicates were performed with a label swap. Samples weredigested with trypsin in a 0.5-mL Amicon Ultra ultrafiltration device (30-kDacutoff; Millipore) (39) with minor modifications. Mass spectrometry wasperformed according to ref. 38. Identification and quantification of peptidespectrum matches were conducted in the framework of Ursgal (40) andusing pyQms (41). See SI Appendix, SI Materials and Methods for details.

Transmission Electron Microscopy. A total of 5 μL of the PSAA–His-Tag purifiedSDG fraction was applied to glow-discharged 400-mesh copper TEM gridscoated with a thin continuous film of evaporated carbon and complexesstained with 2% uranyl acetate (wt/vol). Single particles were imaged on a

Tecnai 12 TEM operated at 120 kV (FEI Company) connected to a DirectElectron LC-1100 lens-coupled 4k × 4k CCD camera (nominal magnification of67,000×, 2× pixel binning, 4.34 Å pixel size at specimen level). An initial 2Dclass average was calculated from manually picked particles to enable auto-mated particle selection using RELION (16). A total of 526,519 particles werecollected from 2,708 digital micrographs (examples shown in SI Appendix, Fig.S4 A and B, circular mask of 380 Å). Single particle images were analyzed withRELION (16) (see SI Appendix, SI Materials and Methods for details).

Spectroscopic Measurements. P700 absorption and electrochromic shift signalmeasurements were performed with a LED pump-probe JTS-10 spectro-photometer (BioLogic) as described previously (10). Single turnover measure-ments used a dye laser emitting at 640 nm, pumped by the second harmonic ofa Minilite II Nd:YAG laser (Continuum). C. reinhardtii cells were harvested andresuspended to a 20 μg·mL−1 chlorophyll concentration [20 mM Hepes, pH 7.2,10% Ficoll (wt/vol)], and incubated (20 min, dark, constant shaking to avoidanaerobiosis). Anaerobic conditions were reached as described above. Toeliminate contribution of linear electron flow, the PSII inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU, 40 μM) was added. Electron flowrates were determined as the product of kox [P700red] (10). More details aboutexperimental procedures are listed in SI Appendix, SI Materials and Methods.

ACKNOWLEDGMENTS. We thank S. Hawat for help with preparation of MS/MS samples and H. Nüsse and U. Keller for their kind permission and assis-tance to glow discharge grids at the EM facility of the Institute for Medicaland Biophysics (University of Münster, Germany). B.H. acknowledges supportfrom Australian Research Council Grants DP130100346 and DP160101018.M.H. acknowledges support from Deutsche Forschungsgemeinschaft (DFG)Grant HI 739/13-1.

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