The structure and function of eukaryotic photosystem I

Biochimica et Biophysica Acta 1807 (2011) 864–877

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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b b a b i o

Review

The structure and function of eukaryotic photosystem I☆ Andreas Busch a, Michael Hippler b,⁎ a b

Department of Plant Biology and Biotechnology, Faculty of Life Sciences, University of Copenhagen, Thorvaldsensvej 40, DK–1871 Frederiksberg C, Denmark Institute of Plant Biochemistry and Biotechnology, University of Münster, Hindenburgplatz 55, 48143 Münster, Germany

a r t i c l e

i n f o

Article history: Received 30 August 2010 Received in revised form 20 September 2010 Accepted 28 September 2010 Available online 2 October 2010 Keywords: Photosystem I Light harvesting LHCI Red algae Green algae Plastocyanin Ferredoxin Electron transfer

a b s t r a c t Eukaryotic photosystem I consists of two functional moieties: the photosystem I core, harboring the components for the light-driven charge separation and the subsequent electron transfer, and the peripheral light-harvesting complex (LHCI). While the photosystem I-core remained highly conserved throughout the evolution, with the exception of the oxidizing side of photosystem I, the LHCI complex shows a high degree of variability in size, subunits composition and bound pigments, which is due to the large variety of different habitats photosynthetic organisms dwell in. Besides summarizing the most current knowledge on the photosystem I-core structure, we will discuss the composition and structure of the LHCI complex from different eukaryotic organisms, both from the red and the green clade. Furthermore, mechanistic insights into electron transfer between the donor and acceptor side of photosystem I and its soluble electron transfer carrier proteins will be given. This article is part of a Special Issue entitled: Regulation of Electron Transport in Chloroplasts. © 2010 Published by Elsevier B.V.

1. Introduction The photosystem I (PSI) from plants is a multi-protein complex that is embedded in the thylakoid membrane of chloroplasts. PSI evolved over 3.5 billion years ago from a simple homodimeric structure into a sophisticated apparatus that consists of a heterodimeric core. It harbors the intrinsic pigments and cofactors for excitation energy and electron transfer and a peripheral antenna providing additional pigments for efficient light-harvesting. As argued earlier [1] this plant PSI is probably the most efficient nano-photochemical machine in nature. A detailed understanding of functional and structural properties of PSI, however, is an absolute prerequisite for its future applications in possible man-made photochemical devises.

2. The structure of eukaryotic Photosystem I The crystal structure of pea PSI and its recent refinements greatly advanced the understanding of the structure and function of eukaryotic PSI [2–4]. The composition of the PSI core and the function of its subunits has been extensively discussed in numerous excellent recent reviews [5–8]. Therefore, we will only present a short summary

☆ This article is part of a Special Issue entitled: Regulation of Electron Transport in Chloroplasts. ⁎ Corresponding author. Tel.: +49 251 83 24790; fax: +49 251 83 28371. E-mail addresses: [email protected] (A. Busch), [email protected] (M. Hippler). 0005-2728/$ – see front matter © 2010 Published by Elsevier B.V. doi:10.1016/j.bbabio.2010.09.009

of the state of knowledge. Readers that are interested in more details should refer to the mentioned reviews. PSI has been shown to be monomeric in plants, green and red algae as well as in diatoms [3,9–13]. It consists of two functional moieties; the core complex (also called reaction center, RC) and the light-harvesting complex (LHCI). The core harbors the components for the light-driven charge separation and the subsequent electron transfer reactions. Furthermore, it binds approximately 100 chlorophylls (Chls) [2] which serve as antenna system to collect light energy. This core antenna is extended by the light-harvesting complex (LHCI) which forms a crescent-shaped structure at the PsaF/PsaJ site of the core and is energetically coupled to the PSI core via the “gap chlorophylls” [3]. Up to now 15 subunits (PsaA to L and PsaN to P) of the eukaryotic PSI core are known and the most recent refinement of the crystal structure of the PSI–LHCI complex to 3.3 Å identified in total 18 protein subunits, 173 Chls, 15 β-carotenes, 3 (4Fe–4S) clusters, and 2 phyloquinones [4]. It is of note that the subunits PsaO and PsaP could not be identified in any plant crystal structure so far [2–4]. PsaA and PsaB form the central heterodimer holding the reaction center P700 and components of the electron transport chain (ETC), A0 (a Chl a), A1 (phylloquinon) and FX (a (4Fe–4S) cluster). Here the charge separation and the initial steps of the ETC occur. Furthermore, the dimer bind about 80 Chls that form the intrinsic light-harvesting antenna [7]. The small stromal subunits PsaC, PsaD and PsaE, which are in close contact to each other, form the binding site for the soluble electron transporter ferredoxin (Fd). PsaC binds the terminal acceptors of the ETC with FA and FB, both (4Fe–4S) clusters. The kinetics of binding and unbinding of Fd will be discussed in more detail in Section 4.2.

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On the lumenal side PsaF and PsaN are important for interaction with the soluble electron carrier plastocyanin (pc) [14,15], which will be discussed in more detail in Section 4.1. Furthermore, PsaF has been shown to be required for the binding of the Lhca1/Lhca4 dimer to the PSI core [3,16] and the crystal structure shows PsaN interacting with Lhca2/Lhca3 suggesting a role in their stabilization [4]. PsaJ has been shown to be required to maintain a proper orientation of PsaF allowing the formation of a proper pc binding site [17,18]. Furthermore, deletion of PsaJ in Nicotiana tabacum lead to an impaired antenna binding and excitation transfer from LHCI to PSI [19]. The subunits PsaH, PsaL, PsaO, PsaP and PsaI have been shown to be important for building the docking platform for LHCII proteins in a process called state transition (discussed in detail in Section 3.4). The different proteins either interact directly with the LHCII proteins or are involved in stabilizing the binding domain ([5] and references therein). PsaG and PsaK are two small membrane proteins that show a 30% sequence homology in Arabidopsis [5]. PsaG is unique to plants and green algae and binds to PsaB and the crystal structure shows PsaG in close contact to Lhca1 [3]. However, deletion of PsaG has no effect on the functional antenna size but PSI content is reduced to 60–80% of wild type level indicating a role of PsaG in stabilizing the PSI complex [20,21]. PsaK binds to PsaA [3] and is involved in Lhca3/Lhca2 binding as revealed by knock-out and gene knock-down studies [20,22,23]. In Chlamydomonas reinhardtii the PsaK/Lhca3 site is a prime side for LHCI antenna remodeling under adverse environmental conditions [24]. It is of note that the most recent structure improvement of plant PSI to 3.3 Å identified an additional density at the PsaK side, but neither PsaO nor PsaP could be modeled into this density [4]. Therefore, it was modeled as a polyadenine and termed it PsaR. It contains one trans-membrane helix and is located between Lhca3 and PsaK, hence a role in LHCI stabilization could be envisioned [4]. However, since no further biochemical data exist for PsaR, its role as PSI subunits remains to be proven. The described subunits can all be found in plant PSI. It is of note that some PSI subunits can be missing in other eukaryotic PSI. For example, a gene coding for PsaP cannot be found in green algae [5] and red algae lack genes encoding PsaG and PsaH [25,26]. A recent detailed genetic analysis of PSI subunits in the red alga Galdieria sulphuraria [26] identified the cyanobacterial PsaM subunit, which is implicated in trimer formation of PSI [27], in the red algal genome. The transitional position of red algae between cyanobacteria and plants is also exemplified by the chimeric nature of the red algal PsaL and PsaF subunits. While in cyanobacteria PsaL is involved in trimer formation of PSI [28], in plants it interacts with the plant specific PsaH to form the LHCII docking site [29]. PsaL of G. sulphuraria does neither contain the conserved sequences required for PSI trimerization in cyanobacteria nor the plant specific sequences for PsaH interaction [26]. As for PsaF of G. sulphuraria, it contains the lysine rich N-terminal sequence that has been shown to be required for efficient pc and cytochrome c6 (cyt c6) binding in plants and green algae [30–32]. It is of note that G. sulphuraria does not contain pc, but uses instead cyt c6 as soluble electron carrier [26]. 2.1. Organization and function of the intrinsic electron transfer chain The reaction center of PSI consists of six Chls organized into two almost identical branches in which each branch is bound by either PsaA (A-branch) or PsaB (B-branch). The central Chl a/a' pair, traditionally called P700, is oriented almost perpendicular to the membrane plane. From there each branch contains two Chl a (accessory Chl and the A0; or ec2A/B and ec3 A/B according to [33] ) and a phylloquinone (A1, PhQA or PhQB). The two branches join again at the 4Fe–4S-cluster Fx, which is followed by two additional 4Fe–4Sclusters FA and FB that are bound to PsaC. The existence of the two almost identical branches raised the question if both sides are active

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for electron transport. Within recent years numerous studies provided solid evidence that both branches participate in electron transfer to Fx [33–39], although the A-branch appears to be the dominating one with an A-branch/B-branch ratio ranging from ~3:2 for green algae to ~ 3–4:1 in cyanobacteria [33,38,40,41]. Despite the faster initial charge separation in the A-branch [33], the subsequent electron transfer to Fx is with ~ 200 ns in the A-branch about ten times slower than the B-branch with ~20 ns [34]. Traditionally the special Chl a pair P700 was considered the site of the initial charge separation. However, recent ultrafast absorption studies demonstrated that the initial charge separation occurs independently in each branch within the accessory Chl a pair (ec2A/B and ec3A/B) [33,42,43]. The authors suggest a model in which the first radical pair would be at the accessory Chls (ec2ec3) either in the A- or B-branch, rather than at the P700. The radical pair ec2+ec3− would then, in a second step, be re-reduced by the P700 [33,43]. The electron is subsequently transferred to the soluble electron acceptor Fd via the 4Fe–4S clusters Fx, FA and FB while P700 is re-reduced by an electron from pc. 3. The eukaryotic light-harvesting antenna associated with PSI 3.1. Core antenna The core antenna of PSI contains in total approximately 100 Chl a and 15 β-carotene of which the majority is bound to the PsaA/PsaB dimer [3]. The pigments in the core antenna are densely packed forming a network for efficient energy transfer to the reaction center. It is of note that the location and orientation of most of the pigments is conserved between cyanobacteria and higher plants even after 1 billion years of evolution [3,44]. In contrast to the cyanobacterial core antenna which contains a species-dependent number of red Chls [45], the majority of red Chls in higher plants and green algae are located in the LHCI (for discussion see Section 3.2.1). While in Chlamydomonas PSI core preparation no red Chls could be observed [46] for Arabidopsis one or two red Chls in the core were proposed [47,48]. However, a more recent study could not identify red Chls in the PSI core [49]. 3.2. Peripheral antenna The peripheral antenna of PSI consists of nuclear encoded chlorophyll binding proteins (Lhca) which are transported into the chloroplast and form a light-harvesting complex (LHCI) which increases the light-harvesting capacity of PSI. LHCs can be found in almost all photosynthetic organisms and are divided according to bound pigments into the Chl a/b binding proteins of green algae and higher plants, the Chl a binding proteins in the red algae and Chl a/c binding proteins of the chromalveolates [50–52]. While the PSI core is highly conserved throughout evolution [3], the LHCI complex shows a high degree of variability in size, subunits composition and bound pigments, which is due to the large variety of different habitats photosynthetic organisms live in. Despite these large differences the supra-molecular structure of the PSI–LHCI complex appears to be conserved with part of the LHCI antenna at PsaF/PsaJ site of the PSI core (see below and Fig. 1). A detailed analysis of the evolution of the different LHC based on their sequence has been done recently [51,52]. Here, we want to give an overview of available biochemical data for the composition of LHCI of different organisms from the red and green lineage. 3.2.1. The green line 3.2.1.1. Higher plants. In land plants six genes (Lhca1–6) encode for the light-harvesting complex proteins [5]. Based on the available crystal structures [2–4] of pea PSI–LHCI complex Lhca1–4 form a crescent

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Fig. 1. Structure and composition of different eukaryotic PSI–LHCI complexes. Schematic models to compare the composition and structure of LHCI complexes of A. thaliana, C. reinhardtii and C. merolae. Shown are the constitutive PSI antennae as discussed in Section 3.2 as well as additional LHCII antenna proteins that associate during state transition as discussed in Section 3.4. Red squares indicate the presence of red chlorophylls in the respective Lhc-protein.

complex at the PsaF/PsaJ site of the PSI core and coordinates 61 Chl in total (Fig. 1). The order of the Lhca proteins in the crystal structures was assigned starting from the PsaG pole of PSI core with Lhca1, Lhca4, Lhca2 and Lhca3 which is in agreement with biochemical and mutagenic studies [29,53,54]. The most recent refined pea crystal structure [4] shows interaction of Lhca1 with PsaG and PsaB while Lhca4 binds to PsaF. Furthermore, Lhca2 is associated with PsaA and PsaJ while Lhca3 binds via PsaA and PsaK to the PCI core. In addition to Chls between the Lhca subunits (termed “linker” Chls), the “gap Chls” are coordinated between the LHCI and the PSI core complex, allowing for efficient energy transfer. The binding of the Lhca proteins is believed to occur in functional dimers. One dimer (also designated as LHCI-730) is formed by Lhca1 and Lhca4 and can be isolated from plants and also reconstituted in vitro [29,55,56]. The second dimer (also designated as LHCI-680), is formed by Lhca2 and Lhca3 [29,57,58]. However, an Lhca2/3 dimer could neither be isolated from plants nor reconstituted in vitro [59]. Additionally spectroscopy studies indicate that Lhca2 and Lhca3 can also form homodimers [60]. Interestingly the antenna system of land plants shows a higher complexity than the crystal structure implies. Using a gel based approach coupled with mass spectrometry in tomato (Lycopersicon esculentum), numerous isoforms for each of the Lhca proteins could be characterized [61]. Furthermore, additional Lhca types like Lhca5 and Lhca6 were identified in Arabidopsis [62]. Both, Lhca5 and Lhca6, are expressed only at a very low level [63] and up to now biochemical data are only available for Lhca5, which was shown to bind in substoichiometric amounts to PSI in Arabidopsis and tomato [61,64]. Cross-linking experiments showed an association of Lhca5 with the LHCI complex, binding to Lhca2 or as a homo-dimer at the Lhca1/4 site if those proteins have been depleted [65]. Furthermore, in vitro reconstitution experiments suggest a hetero-dimer formation of Lhca1/Lhca5 [66], which is supported by the findings that Lhca5 can structurally replace Lhca4 to a small extent in Lhca4 knockout lines [54]. However, so far no precise function for Lhca5 could be identified since knockout plants show no obvious phenotype under standard laboratory conditions [64]. Yet, recently Lhac5 and Lhca6 have been implicated in NAD(P)H dehydrogenase-PSI complex formation, which involved in cyclic electron transfer [67]. The modular layout of the LHCI suggests a rather flexible structure. This idea has been supported by the notion that PSI in different thylakoid compartments vary in their Lhca composition [68] and the differences in Lhca composition of plants grown under different light

conditions [69,70]. However, a more recent study indicated that the stoichiometry of the Lhca proteins are unaffected by different growth conditions [71]. Additionally numerous studies using knockout plants for each Lhca protein clearly showed that each of the Lhca protein has a unique binding place and cannot substituted by another Lhca [53,54,72,73]. A recent electron microscopy study of PSI–LHCI complexes from different Lhca knock-out plants further strengthens the idea of a very rigid LHCI, since the absence of one Lhca leaves a hole in the structure which cannot be filled with another Lhca [54]. However, Lhca5 appears to be an exception since it can replace Lhca4 to a small extent, leading to a stable PSI–LHCI complex in which the supramolecular organization is identical to the wild type [54]. Additionally Lhca4 is apparently not as strictly required for a stable association of the other Lhca proteins as suggested previously [72,73], since Lhca2/3 can associate with PSI in the absence of Lhca4 [54]. The idea that each Lhca has a specific binding site is also supported by the most recent PSI–LHCI crystal structure [4], which refined the structure of the PSI LHCI contact sites. The presence of specific gap Chls and the asymmetric binding site of the core complex impose structural and functional constraints to the binding of the LHCI [4]. Recently a study demonstrated a dual function of the LHCI in Arabidopsis thaliana [74]. Besides light-harvesting, an intact LHCI serves as safety valve for PSI under high-light preventing oxidative damage to the PSI core. It has been proposed that this ability of LHCI might be due to the presence of so called red chlorophylls in the LHCI. “Red Chls” absorb light at a wavelength above 700 nm, implying an uphill energy transfer to the P700 reaction center [75]. Depending on the species those Chls can be found in the PSI core and/or in the LHCI complex [5]. In A. thaliana, one of the best studies models in this respect, the most red-shifted Chls are present in the LHCI, emitting at 735 nm [76]. In vitro reconstitution studies as well as mutant plants lacking different Lhca were used localize the red Chls. Lhca4 harbors the most red-shifted Chl [55,77] as well as Lhca3 which emits at 728 nm [53,59,60]. But also in Lhca1 and Lhca2 red Chls have been identified, emitting at 701–2 nm [5,78]. Extensive mutagenesis studies [78–82] showed that the red Chls can be found at sites A5 and B6 (nomenclature according to [83]) in all four Lhca proteins in A. thaliana. Lhca3 and 4, which harbor the most red-shifted Chls, an asparagine (N) as ligand at A5 can be found while Lhca1 and 2 harbor a histidine (H) residue at this position. Changing the N-residue to a H-residue at position A5 abolished the red-shifted emission in Lhca3 and 4 while the mutation (H)A5(N) in Lhca1 lead to an 11 nm shift towards the red [80]. However, asparagine as ligand

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does not necessarily lead to a red Chl form as the A2 site of LHCII proteins Lhcb1, 2, 3 and 5 has N-residues but the Chls show no red shift [5]. The biological function of these red Chls is still a matter of debate. It has been proposed that they are advantageous under a dense canopy [84]. But also a photo-protective function has been attributed to the red Chls. Due to their low energy level excitation energy is focused in the antenna in domains with close proximity to carotenoids with high efficiency in Chl triplet quenching [85] which leads to efficient dissipation of excess energy.

very similar, with the highest similarity to A. thaliana Lhca2 [91]. Although the LHCI of C. reinhardtii is about 1.5 times larger than in A. thaliana, only 28% of the PSI antenna in C. reinhardtii is constituted by Lhca proteins harboring red Chls compared to almost 50% in A. thaliana [89]. The extent of the red shift is clearly less with Lhca2 harboring the most red-shifted Chl emitting at 712 nm and Lhca4 as well as Lhca9 with Chls emitting above 707 nm [91]. This goes along with the fact that only Lhca2, 4 and 9 have an asparagine residue at the Chl binding site A5 [89,91], which has been shown to be important for the red shift [80].

3.2.1.2. Chlamydomonas reinhardtii. In contrast to higher plants C. reinhardtii encodes nine different Lhca proteins (designated Lhca1 through 9) which are all expressed on protein level [86]. Biochemical analysis of isolated LHCI complex lead to the estimation that it contains between 10 to 18 copies of Lhca proteins which makes the Chlamydomonas LHCI complex significantly larger than its plant counterpart [87,88] (Fig. 1). The presence of a larger LHCI compared to higher plants has been further supported by results of electron microscopy studies [12,13]. It became evident that the main portion of the Lhca subunits forms a crescent around PSI at the PsaF/PsaJ side but additional Lhca monomers may bind to the PsaH side [12] or forming a second layer of Lhca proteins at the PsaF/PsaJ site [13] adding up to a total of 11 to 14 Lhca subunits per PSI core. Recently, a study using isotope dilution mass spectrometry estimated the LHCI to consist of 7.5 ± 1.4 Lhca subunits per PSI [89]. Furthermore, it became evident that the LHCI is quite heterogeneous since several Lhca proteins are not present in a 1:1 stoichiometry with PSI. While Lhca1, 4 and 7 are in a 1:1 ratio, Lhca2, 3, 8 and 9 are only present in a sub-stoichiometric ratio with PSI. For Lhca3 it appears that it is present in an almost 2:1 ratio with PSI. However, the exact composition of the different Lhca proteins may vary under different environmental stresses giving Chlamydomonas a high degree of flexibility [89]. Besides the larger variety of Lhca, the assembly of the LHCI also differs compared to Arabidopsis. In Chlamydomonas an oligomeric complex containing Lhca proteins except Lhca2, 3 and 9 can be isolated from strains lacking the PSI core which implies that the LHCI assembles independently from the PSI core [90]. Recently, Mozzo and coworkers studied the functional properties for each of the nine Lhca proteins showing that despite the low sequence identity between the different Lhca proteins, the pigment binding and spectroscopic properties are

3.2.1.3. Ostreococcus tauri. Prasinophyceae represent a particularly interesting group to study structure and function of light-harvesting proteins since they represent one of the most basal groups in the green lineage [92]. The two best studied organisms, in respect to their LHC composition, are Mantionella squamata and the recently fully sequenced O. tauri [93]. Their LHC system is characterized by the presence of prasinophyte-specific (Lhcp) antenna proteins that are only distantly related to LHCI and LHCII [52,94] as well as the binding of Chl a, b and c as light-harvesting pigments [94]. Unlike the photosystem-specific association as seen in higher plants or other green algae, only one functional antenna system has been described in M. squamata serving as LHC for both photosystems [95,96]. A surprising complexity of different light-harvesting proteins has been reported for O. tauri (Table 1), encoding for six different LHCI proteins, two classes of Lhcp as well as two minor LHCII-family proteins annotated as CP26 and CP29 [52,94]. While association of Lhca2 with PSI has been shown previously [94], the localization of the other lightharvesting proteins remained unclear. Recently a detailed study of the PSI antenna of O. tauri revealed the binding of all LHCI proteins, the Lhcp as well as the two minor LHCII proteins to PSI [97]. It is of note that the LHCI in O. tauri does not contain far red Chls [97]. 3.2.1.4. Physcomitrella patens. In contrast to higher plants and green algae, little work has been done on the moss PSI and its antenna system complex, although they pose an interesting step in evolution, representing one of the first steps in the transition from aquatic to terrestrial life [98]. Recently, a detailed in silico and biochemical study of the PSI–LHCI complex of Physcomitrella patens was published [99]. The authors used the available EST collections and the fully sequenced genome [98] to identify light-harvesting proteins and performed a

Table 1 Overview of identified genes encoding for chlorophyll binding light-harvesting genes potentially interacting with PSI. Known interactions are discussed throughout the text. Organism Green clade Arabidopsis thaliana* Physcomitrella patens*

Ostreococcus tauri*

Chlamydomonas reinhardtii* Red clade Cyanidioschyzon merolae* Porphyridium cruentum Galdieria sulphuraria (also designated Cyanidium caldarum) Phaeodactylum triconutum*

Cyclotella cryptica

* indicates organisms with sequenced genome.

Gene names

References

Atlhca1, Atlhca2, Atlhca3, Atlhca4, Atlhca5, Atlhca6 Pplhca1.1, 1.2, 1.3 Pplhca2.1, 2.2, 2.3, 2.4 Pplhca3.1, 3.2, 3.3, 3.4 Pplhca5 Otlhca1, Otlhca2, Otlhca3, Otlhca4, Otlhca5, Otlhca6 Otlhcp1, Otlhcp2.1, Otlhcp2.2, Otlhcp2.3, Otlhcp2.4 CP26, CP29 Crlhca1, Crlhca2, Crlhca3, Crlhca4, Crlhca5, Crlhca6, Crlhca7, Crlhca8, Crlhca9

[5] [99]

CMN235C, CMN234C, CMQ142C PclhcR1,PclhcR2 Gslhcr1, Gslhcr2, Gslhcr3, Gslhcr4, Gslhcr5 Designation according to [107] Group1 (Lhcf): fcpA, fcpB, fcpC, fcpD, fcpE,fcpF Designation according to [110] Group1 (Lhcf): lhcf1 through 17 Group2 (Lhcr-like): lhcr1 through 14 Group3 (LhcSR-like): lhcx1 through 4 Group1(Lhcf): fcp1, fcp2, fcp3, fcp4 Group2 (Lhcr-like):fcp4 Group3 (LhcSR-like): fcp6, fcp7, fcp12

[25] [103,236] [104,105] [107,110]

[52,94,97]

[86]

[108,237]

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comprehensive homology analysis between moss, plant and algal Lhc protein sequences. It became evident that P. patens encodes more Lhc proteins than A. thaliana (Table 1), but no A. thaliana Lhca4 and Lhca6 homolog could be identified [99]. This is of special interest since Lhca4 harbors the most red-shifted Chl in A. thaliana, with an emission of 735 nm [80]. This lack of Lhca4 homologs in P. patens is reflected in the less red-shifted 77 K fluorescence emission spectra of isolated PSI– LCHI complex [99]. Biochemical characterization of isolated PSI–LCHI complex from P. patens using native gel electrophoresis revealed that the complex is slightly smaller compared to its Arabidopsis counterpart. The authors suggested that this might be due to the presence of only three light-harvesting proteins in the complex. However, this needs to be analyzed in more detail and appears to be highly unlikely in the light of the different structural data from green and red algae LHCI that all showed a crescent-shaped LHCI at the PsaF/PsaJ site, which can harbor at least four Lhca proteins [10–13]. It is of note that for the red algae Cyanidioschyzon merolae, which encodes only three Lhca proteins, a crescent-shaped Lhca belt as known from plant PSI– LHCI has been shown [10]. 3.2.2. The red line While red algae contain a primary plastids, secondary endosymbiosis of a red algae by another eukaryote led to a huge variety of algae in the red line, including haptophytes, cryptophytes, stramenopiles (including e.g. diatoms), dinoflagellates and apicomplexans [100]. While it is generally accepted that these groups have red algal derived chloroplasts, the sequence and number of endosymbiotic events is still debated (see e.g. [100] and references therein). With the increasing number of sequence information for different organism more and more focus is put on the actual biochemical characterization of PSI–LCHI. Here we will discuss some examples for which a reasonable amount of biochemical data are available. 3.2.2.1. Red algae. While retaining phycobilisomes for light-harvesting for PSII, rhodophyta possess light-harvesting proteins (termed Lhcr) for PSI which bind Chl a [101]. The Lhcr proteins have been shown to be exclusively associated with PSI [10,101,102] and their number is species-dependent. Biochemical data showed six different Lhcr in Porphyridium cruentum [103], five in Galdieria sulphuraria [104,105] and three in C. merolae [10]. The arrangement of Lhcr proteins at the PSI-core was investigated by electron microscopy using isolated PSI particles from Cyanidium caldarium [11] and C. merolae [10]. Overlaying the crystal structure from pea showed that the organization of the LHCI follows the same principle as in higher plants with the Lhc proteins forming a crescent at the PsaF/PsaJ site [10,11]. However, studying the PSI associated antenna system of C. merolae in more detail it became evident that although C. merolae only encodes three different Lhcr [25], a crescent of four Lhcr at the PsaF/PsaJ site is formed, indicating that one Lhcr is present in an at least 2:1 stoichiometry (Fig. 1) [10]. Interestingly, a PSI specific phycobilisome antenna capable of transferring energy to the PSI core could be detected (Fig. 1) [10]. However, the precise composition and function of this phycobilisome form remains to be elucidated. 3.2.2.2. Diatoms. Diatoms are unicellular eukaryotic algae that can be found in oceans, freshwater and in soil and are responsible for almost one quarter of global primary production [106]. The pigmentation of the light-harvesting complexes differs greatly from higher plants and green algae. The fucoxanthin-chlorophyll-proteins (FCPs), which serve as light-harvesting complexes, bind Chl c instead of Chl b and fucoxanthin instead of lutein. Despite these differences FCPs can be placed into the group of Chl a binding proteins [107,108]. Based on phylogenetic analysis [51,52] FCPs are divided into three clades (Table 1): (i, Lhcf) FCPs that are considered to be involved in lightharvesting and group with the most commonly isolated FCPs from Chl c containing algae. Furthermore, (ii, Lhcr-like) FCPs that group with

Chl a containing red algal Lhcr and (iii, LhcSR-like or Lhcx) those FCPs that group with green algal LhcSR (Li818) genes which have been implicated in protection against high-light [109]. Two of the most studied diatoms in respect to the biochemistry of FCPs and their organizations are the centric diatom Cyclotella meneghiniana, the closely related Cyclotella cryptica and the pennate diatom Phaeodactylum triconutum of which the genome was sequenced recently [110]. In C. meneghiniana different oligomeric structures of FCPs have been isolated and the associated FCPs polypeptides were identified [111,112]. A trimeric structure, termed FCPa, is mainly composed of 18 kDa subunits encoded by fcp2, but also a 19 kDa subunit, encoded by fcp6, could be detected in smaller amounts. Furthermore, hexa- or nonamers (FCPb) could be detected. Those are composed of only 19 kDa subunits most likely encoded by fcp5. Also in P. tricornutum FCP trimers or hexamers, with subunits encoded by fcpC/D/E can be found [113,114]. A more detailed study using His-tagged FcpA revealed that the FCP trimer is the basic building block of FCPs and is built up by FcpA and FcpE [115]. However, the specific stoichiometry of the two subunits remains to be elucidated. A possible specific association of FCPs to one of the photosystems is still controversial. Unlike in plants the thylakoids are not organized in grana and stroma lamella but in bands consisting of three thylakoids each [116] in which PSI and PSII are randomly distributed [117]. Berkaloff and co-workers concluded that FCPs serve as antenna for both photosystems [118] which is in agreement with work done in C. cryptica where in both PSI and PSII fractions Fcp2 and Fcp4 proteins could be found [119]. However, more recent work, using more gentle solubilization methods, could demonstrate a specific FCP binding to PSI both in P. tricornutum [9] and C. meneghiniana were a PSI specific antenna is formed by Fcp5, a Fcp4-like Fcp and a 17 kDa Fcp [120]. 3.2.2.3. Dinoflagellates. Dinoflagellates contain two types of lightharvesting complexes [121]. The membrane bound LHC binds Chl a/c and peridinin as carotenoid and is related to the FCPs of chromophytes. Interestingly these proteins are encoded as polyproteins which are imported into the chloroplast where they are cleaved into numerous mature proteins [122,123]. Additionally a unique soluble light-harvesting system, the peridinin-chlorophyll a protein (PCP), which is unlike any other known protein [124] increases the lightharvesting capacity. PCP is located in the lumen of the chloroplasts and uses carotenoids as main light-harvesting pigment instead of Chls, allowing absorption of more blue-green light (470–550 nm) [125]. The structure of the PCP of the dinoflagellate Amphidinium carterae is the best-studied PCP and has been resolved to 2 Å resolution [126] providing valuable insight to understand the energy transfer within the PCP. The structure of the main form of PCP (MFPCP) consists of a protein of 313 amino acids forming two pseudosymmetrical α-helices. Each helix holds four peridinins arranged in van der Waals contact around a central Chl a molecule. Via hydrophobic interaction, PCP monomers form trimers with a diameter of about 100 Å and a thickness of about 40 Å [126]. So far only one other PCP form has been reported, the high-salt PCP form (HSPCP). HSPCP constitutes about 2% of all PCP in A. carterae and is named due to the fact that this PCP form elutes at higher salt concentrations from cation exchange column than the MFPCP [127]. HSPCP shows only a 31% sequence homology but also forms a twosubunit domain structure. However, HSPCP bind only six peridinins instead of eight like MFPCP. Recently the structure of HSPCP has been resolved to 2.1 Å [128] demonstrating a drastic difference of pigment and lipid orientation compared to MFPCP. PCP can be reconstituted from heterologously expressed apoprotein and purified pigments [129] which also allows the substitution of pigments of interest to study binding affinities and specificities [130]. This makes PCP an ideal model to study energy transfer from carotenoids to Chl. Hence PCP have been subject to numerous spectroscopic studies to reveal the

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energy transfer within the protein complex and the obtained results have been reviewed in detail recently [131]. 3.3. Energy transfer between the peripheral and core antenna One of the major determinant for efficient excitation energy transfer between two pigments is the distance between the two pigments, according to the Förster approach to the power of 6 [132]. Therefore the discovery of the gap Chls between the PSI core and the LHCI was a key to model the kinetics of energy transfer [3]. The crystal structure revealed 10 gap Chls which are located in the space between PSI core and LHCI [3]. However, the gap Chls are distributed unevenly with seven of the pigments located close to the Lhca1/Lhca4 dimer, indicating a preferred excitation energy transfer route. The most recent refinement of the PSI-crystal structure identified additional gap Chls between PsaB and Lhca4/Lhca1 as well as one being coordinated by Lhca2 allowing energy transfer from Lhca2 to the PSI core [4]. Additionally, a gap carotenoid was identified in the space between PsaA and Lhca2/Lhca3. These additional gap pigments complemented the picture for potential excitation energy transfer routes from LHCI to the PSI core. The importance of the gap Chls has been demonstrated in a computational modeling approach simulating excitation energy transfer networks in PSI. Indeed without gap Chls the LHCI to PSI core energy transfer times rises from 2.1 to 10.5 ps [133]. Excitation energy transfer has been studied in Lhca1 and Lhca4. After excitation of Chl b the energy rapidly moves to neighboring Chls in the time scale of 0.2–0.7 ps, while the equilibration with other the Chls, including the terminal emitter, occurs within 3–5 ps [134,135]. Furthermore, an excitation energy transfer with a lifetime of 15–30 ps could be observed in Lhca1/Lhca4 dimers, which was attributed to an inter-subunit energy equilibration [136]. Time resolved spectroscopy measurements of isolated PSI–LCHI complexes from Arabidopsis [75,137,138] and Chlamydomonas [139,140] revealed a photochemical trapping time in the PSI core antenna of 20–50 ps and a second slower excitation trapping phase of 70–150 ps which was assigned to the LHCI to PSI core energy transfer. This slow LCHI to PSI core energy transfer is rather surprising since the existence of gap Chls suggested an efficient and fast energy transfer and computational kinetic calculated energy transfer rates between 1.5 to 6 ps [133]. It has been proposed that the slow energy equilibration between LHCI and PSI core is due to the existence of the red Chls in the LHCI antenna, which serve as excitation sinks, especially at low temperatures [139]. This idea has been supported by trapping kinetics of isolated PSI–LHCI complexes of Arabidopsis and Chlamydomonas [47]. Although the LHCI of Chlamydomonas is about 1.5 times bigger than its Arabidopsis counterpart the energy trapping time was faster in Chlamydomonas than in Arabidopsis. These differences were attributed to the red Chls of which Arabidopsis a larger proportion in its LHCI (as discussed in 3.2.1). 3.4. State transitions To balance the excitation energy between PSI and PSII, photosynthetic organisms evolved a mechanism termed state transitions [141– 143]. Under light conditions which preferentially excite PSII, both PSII core and LHCII proteins become phosphorylated [144], LHCII proteins then dissociate from PSII and move to PSI (state 2). This process is reversible, when PSI is more excited PSII core and LHCII proteins become de-phosphorylated [70] and migrate back to PSII (state 1). The extent of moveable PSII antenna differs significantly throughout different taxa. While in the green algae C. reinhardtii the proportion of displaced LHCII antenna is about 80% [145], in higher plants such as A. thaliana only 15–20% [146] of LHCII is mobile and in Prasinophytes such as O. tauri no state transition could be observed [97]. Despite this large difference in the extent of movable antenna it is believed that the mechanistic basis of state transition is similar in all LHCIIcontaining organisms [147]. A central role in state transition plays the

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kinase STT7 that was shown to be essential for LHCII phosphorylation in Chlamydomonas [148]. In Arabidopsis two homologs of STT7 could be identified, STN7 and STN8 [144,149]. While STN8 is required for phosphorylation of several PSII core subunits [144,150], STN7 is a functional homolog of STT7 [149]. For a recent review on the structure and function of STT7 and its homologues the reader is referred to [151]. On the PSI side, the subunits PsaH, PsaL and PsaO form the docking site for phosphorylated LHCII (pLHCII), which has been demonstrated using cross-linking experiments and RNA interference approaches [152–154]. Hence, Arabidopsis plants lacking PsaO are reduced 50% in state transition [154], while plants lacking PsaH or PsaL are locked in state 1 [152]. As for the movable LHCII antenna, using electron microscopy and image analysis of digitonin solubilized PSI in state II from Arabidopsis, a large density at the PsaH/L/A/K side could be identified, which was assigned to a LHCII trimer (Fig. 1) [155]. More details of the movable LHCII antenna are known for Chlamydomonas. Takahashi and co-workers [156] were able to isolate a state 2-specific PSI–LHCI supercomplex which harbors two minor monomeric LHCII proteins CP26 and CP29 as well as a major LHCII protein designated Lhcbm5 (Fig. 1). Since Lhcbm5 has no homolog in higher plants and show similarity to CP26 and CP29, the authors suggested a CP24-like role of Lhcbm5 in Chlamydomonas [156]. RNAi strains with reduced CP26 or CP29 level showed normal LHCII phosphorylation [157]. However, while the CP26 RNAi line had a normal association of pLHCII to the PSI complex, the CP29 RNAi lines completely lacked pLCHII association to PSI [157]. This indicates the central role of CP29 in the LCHII-PSI attachment process in Chlamydomonas. Upon induction of state 2, CP29 becomes hyperphosphorylated, detaches from PSII and migrates to PSI where it binds to the PsaH site and serves as a docking site for further LHCII proteins [158]. Using fluorescence lifetime imaging microscopy (FLIM) Iwai and co-workers [159] visualized the detachment of pLHCII from PSII during state transition in vivo. Upon induction of state 2, pLHCII aggregates accumulate in the thylakoid membranes from which only a part associate with PSI. It was suggested that the remaining free pLHCII aggregates can serve as an efficient quencher protecting the cells from photo-oxidative damage. 4. Electron transfer between the donor and acceptor side of PSI and its soluble electron transfer carrier proteins It is instructive to note that the study of photosynthetic intermolecular electron transfer reactions offers the means for the investigation of protein–protein-interaction at high resolution because the binding and electron transfer steps can be readily explored spectroscopically. These measurements provide kinetic and binding constants that can be used to deduce the mode and mechanism of the protein–protein-interaction (for recent reviews see [160–163]). Intermolecular electron transfer reactions are important not only in photosynthesis but also in many biological processes including respiration. In these reactions, two electron transfer proteins bind to each other to form an electron–donor–acceptor complex. Recent progress in understanding the dynamics of recognition and binding in such complexes indicates the interplay of two types of interaction. The first, the long-range interactions, guided by electrostatic forces serve to orient both partners. The second, based on hydrophobic interactions, serve to optimize the orientation of both partners to permit efficient short-range electron transfer (exemplified by publications [30,31,161,163–174]). 4.1. Donor side of PSI Binding and electron transfer between pc or cyt c6 and PSI can be described by the outlined two-step model. In eukaryotic organisms, binding of pc or cyt c6 to the PSI is mainly driven by electrostatic

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attraction and hydrophobic contact (Fig. 2). Long range electrostatic attractions between basic patches of PsaF and acidic regions of pc and cyt c6 [30,31,175–177] as well as the hydrophobic contact between the electron transfer site of the donors and of PSI including PsaAW651 and PsaB-W627 in C. reinhardtii [171,176] are required for stable electron transfer complex formation and efficient electron transfer. 4.1.1. Requirement of PsaF Studies using chemical cross-linking and mass spectrometry as well as knock-out and reverse genetics experiments have established the crucial function of the positively charged eukaryotic N-terminal domain of PsaF for the binding of both donors [15,16,30–32,178]. It was found that Lys23 of PsaF from C. reinhardtii is the key residue for the electrostatic interaction between PSI and pc or cyt c6 [30,31,178]. Interestingly, the positively charged N-terminal domain is missing in prokaryotic organisms and, in line with this finding, efficient binding and electron transfer between PSI and its donors pc or cyt c6 are independent of PsaF, as revealed by knock-out experiments and functional electron transfer measurements in the cyanobacterium Synechocystis sp. PCC6803 [30,179,180]. On the other hand, introduction of the basic PsaF domain from C. reinhardtii into cyanobacterial PsaF allowed efficient binding and fast electron transfer between the algal donors and the chimeric cyanobacterial PSI. This proves that the eukaryotic positively charged N-terminal domain of PsaF is instructive and essential for electrostatic attraction of the donors and formation for the electron transfer complex competent in fast electron transfer [32]. 4.1.2. The lumenal loops i/j of PsaA/B form a hydrophobic recognition site for binding of pc and cyt c6 to PSI In contrast to the N-terminal domain of PsaF, the hydrophobic interaction site of the PSI core formed by PsaA and PsaB important for efficient docking of the electron transfer donors is conserved between

prokaryotic and eukaryotic organisms. This site is provided by luminal loops i/j of PsaA and PsaB [171,172,181], where residues PsaA-W651 and PsaB-W627 constitute the key elements, having their indole groups stacked at van der Waals distance [44] and being directly situated above P700 [3,44] (Fig. 2). The importance of the luminal loop j of the PsaB for electron transfer between PSI and the soluble donors has been first demonstrated in Synechocystis PCC 6803. The introduction of a double mutation in the loop j of the PsaB (W622C/A623R) led to a highly photosensitive phenotype and caused a severe defect in the interaction and electron transfer with pc or cyt c6 [181]. Further experiments addressed the specific function of the two Trp residues forming the sandwich-like structure above P700. The change of either of the two PsaA or PsaB Trp residues to Phe in C. reinhardtii abolished the formation of an intermolecular electron transfer complex between the altered PSI and pc, indicating that PsaATrp651 as well as PsaBTrp627 of loops i/j form the hydrophobic recognition site required for binding of pc to the core of PSI [171,172]. 4.1.3. Stabilization and orientation of the N-terminal domain of PsaF and refinement of binding and release of both donors in eukaryotic organisms The luminal loop j of PsaB has, besides its importance for docking of the donors, additional functions related to orientation of the Nterminal domain of PsaF and unbinding of oxidized pc. Site-directed mutagenesis of PsaB-Glu613 to Lys resulted in a strong impairment of electron transfer between PSI and the two donor proteins as investigated by in vitro flash-induced absorption spectroscopy [171]. The electron transfer rates for the mutant Glu613Lys PSI and the donor pc or cyt c6 were comparable to values obtained for electron transfer between PSI from the PsaF-deficient mutant and the two donors. In contrast, another mutation of PsaB-Glu613 to Asn had the opposite effect. It resulted in a clear acceleration of electron transfer rates and an improved binding of both donors to the altered PSI in respect to wild type. The structural data clearly show that the negative

Fig. 2. Binding of plastocyanin (pc) to the oxidizing site of PSI. Long range electrostatic attractions between basic patches of PsaF and acidic regions of pc are guiding pc into the hydrophobic binding pocket of PSI mainly formed by PsaA-W651 and PsaB-W627 allowing close contact between the copper (blue dot) and His87 of pc with P700 for fast and efficient electron transfer [2,172,178,235]. PsaB-residues Asp612 and Glu613 are essential for efficient unbinding of oxidized pc. Red shows negatively and blue positively charged amino acid residues. The bold parts in PsaF subunits represent Lys residues of the N-terminal domain.

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charge provided by residue PsaB-Glu613 is exposed on the surface of PSI [3]. Thus, it is possible that a change of Glu to Asn leads to a tighter binding of the donors to PSI in mutant PsaB-Glu613Asn since this mutation decreases the electrostatic repulsion between the negatively charged donors and PSI, which in turn might lead to a slower unbinding of oxidized pc. Equally, PsaB-Glu613 of C. reinhardtii is at close distance to the N-terminus of the PsaF subunit of the plant PSI and might therefore be important for the orientation of the positively charged N-terminal domain of PsaF in C. reinhardtii. In line with this consideration, the mutation of PsaB-Glu613 to Lys indeed caused a distortion of that domain by charge repulsion and led to a disorientation of the N-terminal domain of PsaF and a loss of affinity for binding of the two donors to PSI as revealed by the measurement of the electron transfer properties for the electron transfer between PSI PsaB-Glu613Lys and its donors (see above). To summarize, residue PsaB-Glu613 might be important for efficient unbinding of oxidized pc, a conclusion confirmed by in vivo electron transfer measurements (see below), as well as for orientation of the positively charged Nterminal domain of PsaF in a way that it allows efficient binding of the donors to PSI [171]. The latter conclusion is strongly supported by the recent resolution structure of Pea PSI [2,182], where PsaB-Glu613 was found to be in close contact to Lys residues of the N-terminal domain of PsaF. 4.1.4. Kinetic analysis of the electron transfer between soluble donors and PSI The electron transfer between pc or cyt c6 and the primary donor of PSI, P700, can be measured after the photooxidation of P700 by a short flash. In plants and green algae, the electron transfer reaction generally displays two distinct kinetic phases [183–186]. A simple explanation for this biphasic kinetic scheme was that PSI particles which have already formed a stable complex with the donor prior to the flash give rise to fast microsecond intra-molecular electron transfer. The remaining PSI complexes are re-reduced by the soluble donor in a bimolecular reaction with second-order kinetics. Evidence for such a stable pre-formed complex was only found in the presence of the eukaryotic positively charged domain of subunit PsaF (see above), where this intra-molecular complex can be also stabilized by chemical crosslinking in a way that the rate of electron transfer between the crosslinked donor and PSI is comparable to that found in the native complex [178,187]. The formation of a stable intramolecular electron transfer complex between PSI and pc also impacts the thermodynamic properties of this electron transfer reaction. It was determined that the dissociation constant of oxidized pc is about six times larger than that observed for reduced pc [188]. Consistent with this result the midpoint redox potential of pc bound to PSI was found to be 50–60 mV higher than that of soluble pc. Although this would lead to a decrease in the driving force of the intra-molecular electron transfer complex, the turn-over of electron transfer between pc and PSI would be optimized due to a faster unbinding of oxidized pc. 4.1.5. Release of oxidized pc from PSI limits linear electron transfer between cytochrome b6/f complex and PSI in eukaryotic organisms Investigation of binding dynamics and electron transfer between pc and PSI, and cytochrome f oxidation kinetics in C. reinhardtii, mutants using fast absorption spectroscopy revealed that the pc, cyt b6f and PSI pools almost approach redox equilibrium in vivo [189]. Remarkably, under these conditions, electron flow between PSI and the cytochrome complex was not affected by a five-fold lower binding affinity of pc to PSI (mutant PsaA-W651Ser,[171]). On the other hand, electron flow from PSI to the cytochrome b6f complex was very sensitive to a two to three-fold decrease in the rate of pc release from PSI (mutant PsaB-Glu613Asn,[171]), suggesting that this is the kinetic step that limits electron transfer between cytochrome b6f complex and PSI in vivo [189]. This results confirmed the notion that the

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mutation of PsaB-Glu613Asn leads to a slower unbinding of oxidized pc (see above) and substantiated experimentally previous modeling of electron transfer at the donor side of PSI that suggested that the release of oxidized pc from PSI might represent the kinetic limiting step in electron transfer between cyt b6f complex and PSI (see above [188]). These results indicate that during evolution binding of reduced pc was optimized in a way that unbinding of oxidized pc limits linear electron transfer between the cytochrome b6/f complex and PSI in chloroplast-based photosynthesis, indicating that fast reduction of photo-oxidized P700+ is of physiological importance for these systems. 4.2. Acceptor side In contrast to the oxidizing side, the reducing side of PSI is well conserved between cyanobacterial and chloroplast-based PSI [3,44]. So consequently data obtained with cyanobacterial systems are highly relevant for the description of electron transfer reaction between Fd and PSI in oxygenic chloroplast-based photosynthesis. At the acceptor side of PSI three peripheral subunits protrude into the stromal space: PsaC, PsaD and PsaE. The three subunits are close together and form a ridge on the stromal side. While subunits PsaD and PsaE do not carry any cofactors, the PsaC subunit binds the terminal electron acceptors FA and FB each harboring one (4Fe–4S) cluster which are coordinated by two sets of four cysteine residues [190,191]. Site-directed mutagenesis of the cysteine ligands in cyanobacteria demonstrated that Cys 11, 14, 17 and 58 coordinate the iron atoms of cluster FB whereas Cys 21, 48, 51 and 58 coordinate cluster FA [192]. Chemical inactivation studies as well as reverse genetics experiments supported the notion that FB represents the distal cluster [193–195] implying that electron transfer in the terminal part of PSI follows the FX-FA-FB-Fd pathway. This view is well supported by the 2.5 Å structure of cyanobacterial PSI [44] and indicates that electron transfer within PsaC takes place against the redox potential gradient as FA is more electropositive than FB [196,197]. 4.2.1. Kinetic analysis of the electron transfer between soluble acceptor and PSI In chloroplast-based photosynthesis, Fd functions as an electron acceptor for PSI. Fd is a soluble iron-sulfur protein harboring a (2Fe– 2S) cluster. This cluster is reduced at the stromal side of PSI by the FB cluster of PsaC. The photoreduction of Fd by cyanobacterial or algal PSI is characterized by first-order components followed by a secondorder electron transfer process [198–201]. The formation of the intermolecular electron transfer complex between Fd and PSI depends mainly on electrostatic interactions, for which subunits PsaC, PsaD and PsaE sign responsible. 4.2.2. Requirement of PsaD, PsaE and PsaC for reduction of Fd The requirement of PsaD, PsaE and PsaC for reduction of Fd has been recently reviewed in detail [202] and will be here addressed briefly. Chemical crosslinking studies first demonstrated a physical interaction between soluble Fd and PsaD [203]. Mass spectrometric analyses of this crosslinked complex obtained from Synechocystis sp. PCC 6803 revealed crosslinking sites between PsaD and Fd [204]. However the site directed change of the Lys residue of PsaD (Lys106) involved in crosslinking with Fd had a relatively small effect on binding and electron transfer to Fd [205,206]. On the other hand, deletion of PsaD has a stronger impact on binding and electron transfer of the PsaD-deficient PSI to Fd [206,207], yet, flash induced absorption spectroscopy indicated that first-order electron transfer components were still present. The impact on the electron transfer reaction was observed as a ten-fold decrease of the second-order rate constant for reduction of Fd by mutant PSI as compared to wild type PSI [208], pointing to the fact that PsaD is crucial for electrostatic interaction but not essential for the formation of the inter-molecular

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electron transfer complex formed between Fd and PSI. PsaD mutants in A. thaliana had impaired photosynthetic electron transfer, suffered from photoinhibition, possessed decreased PSI stability and altered expression of nuclear genes for chloroplast functions [209,210]. It is of note that the deletion of both PsaD alleles encoded in the A. thaliana genome is seedling-lethal, pointing to an essential role of PsaD for PSI stability and/or PSI function in vascular plants. Deletion of PsaE increased the dissociation constant for Fd binding about 50-fold [208], whereas second-order electron transfer rate was almost unaffected. These data strongly suggested that PsaE is involved in the stabilization of the inter-molecular electron transfer complex. Further reverse genetic experiments revealed PsaE-Arg39 from Synechocystis sp. PCC 6803 to be a key residue for binding and stabilizing Fd in the electron transfer complex with PSI [211,212]. Interestingly an Arabidopsis mutant lacking psae1-3 and psae2-1 genes and therefore devoid of the PsaE protein was still able to grow photoautotrophically, implying that PsaE is not essential for PSI accumulation and thylakoid electron flow in vascular plants [213]. Work conducted with C. reinhardtii demonstrated that the PsaC subunit of PSI possesses a central role in the binding of Fd to the stromal side of PSI [198]. In particular reverse genetics recognized PsaC-Lys35 as a key interaction site for contact between PSI and Fd since site-directed mutations of Lys to uncharged or negatively charged amino acids abolished the formation of a firstorder electron transfer complex. Furthermore, alteration of PsaCLys35 by site-directed mutagenesis eliminated chemical crosslinking between Fd and PsaD, PsaE, and PsaC. Thus PsaC, in particular PsaCLys35, is essential for the formation of the inter-molecular electron transfer complex between PSI and Fd. In conclusion, PsaC, PsaE and PsaD contribute differently to binding and complex stabilization, (i) PsaD appears to be important for the electrostatic guidance of Fd into the PSI binding pocket, (ii) PsaE, and in particular PsaE-Arg39, have a role in stabilizing the final electron transfer complex, while (iii) PsaC via Lys35 establishes close protein contact required for fast electron transfer between the ironsulfur cluster of Fd and the terminal electron acceptors (FA; FB) of PSI. 5. Structure and function of PSI in cyclic electron transfer A crossroad in photosynthetic electron transfer is at the level of reducing power generated by PSI. These “highly reducing” electrons can either drive linear electron flow (LEF) or cyclic electron flow (CEF). In CEF electrons from the reducing side of PSI are re-injected into the photosynthetic electron transport chain either at the level of the plastoquinone pool or at the stromal side of the cyt b6 /f complex. CEF around PSI has been first recognized by Arnon [214] and has attained much attention in the recent years [215–220]. CEF is considered to have an important function in the re-equilibration of the ATP poise and avoidance of over-reduction of the PSI acceptor side [215–220]. In microalgae and vascular plants, CEF is reliant on NAD(P) H dehydrogenase (NDH)-dependent and/or PROTON GRADIENT REGULATION5 (PGR5)-related pathways [215,218,220]. For both pathways, supercomplexes consisting of PSI–LHCI and components of the respective electron transfer routes have been recognized in vascular plants. In A. thaliana a novel NDH-PSI supercomplex with a molecular mass of N1000 kD was discovered [221]. In mutants lacking NdhL or NdhM, a putative subsupercomplex with a slightly lower molecular mass was found consistently [221]. Interestingly, the minor LHCI-subunits Lhca5 and Lhca6 appear to be required for the formation of this full-size NDH-PSI supercomplex [67]. There is further evidence that the interaction of NDH and PSI advances the in vivo function of NDH [67]. The expression of Lhca5 on protein level is rather low and for Lhca6 no endogenous expression at protein level has been confirmed (see above). Therefore the formation of the NDHPSI supercomplex is clearly substoichiometric, indicating that the other CEF pathway could operate at the same time. In C. reinhardtii, the non-photochemical reduction of the PQ pool does not involve the

NDH-1 complex, which is missing from the algal chloroplast, but instead a type II NADH dehydrogenase called Nda2 [222–224] functions as a PQ-reductase. It is of note that so far no evidence for functional supercomplex formation between Nda2 and PSI has been reported. For the PGR5-dependent CEF pathway [225], the thylakoid protein PGR5-Like 1 (PGRL1) was discovered as a novel component required for CEF in Arabidopsis [226]. It is suggested that PGRL1 and PGR5 interact physically in A. thaliana, and that this complex associates with PSI, thereby facilitating the operation of PSI cyclic electron transport [226]. In the green alga C. reinhardtii, where CEF is largely induced under state 2 conditions (as explained in more detail above), Iwai et al. [227] isolated from these conditions a protein supercomplex composed of PSI–LHCI, LHCII, the cytochrome b6f complex, Fd (Fd)-NADPH oxidoreductase (FNR), and PGRL1. Functional spectroscopic analyses indicated that upon illumination, electrons from the reducing side of PSI were transferred to Cyt bf via soluble Fd. Additionally in the presence of pc, the complex showed re-reduction of oxidized PSI and oxidation Cyt f. In conclusion, this supercomplex appears to be capable of CEF [227] by shuttling electrons through the complex via the support of the soluble electron transfer proteins pc and Fd. This sheds further light on the mechanistic properties of CEF, namely that CEF can operate at the reducing side of PSI via electron transfer between Fd and Cyt b of cytochrome b6f complex as suggested earlier [228]. It is possible that FNR, which was previously found to be associated with the cytochrome b6f complex [229], also participates in Cyt b photoreduction. As mentioned above PGRL1 was identified in the algal supercomplex. In C. reinhardtii PGRL1 was found to be induced under low iron conditions and necessary for efficient CEF under iron-deprivation [230], therefore indicating a role of PGRL1 in CEF as described for A. thaliana [226]. At the same time there is experimental evidence that PGRL1 participates in ironinduced remodeling of the photosynthetic apparatus [230]. It is interesting to note that PGR5, that was not identified in the algal CEFsupercomplex [227], is also strongly induced at the mRNA level under iron-deficiency in C. reinhardtii. It is possible that PGR5 and PGRL1 operate both in iron homeostasis and CEF. Thiol-trapping experiments indicated iron-dependent and redox-induced conformational changes in PGRL1 [230] that may provide a link between iron metabolism and the partitioning of photosynthetic electron transfer between linear and cyclic flow. Another level of regulation could be introduced via the expression of distinct Fd isoforms that are reduced by PSI, but possess different binding and electron transfer activities towards components of LEF and CEF. Such a partitioning has been first observed with two Fd isoforms from the C(4) plant maize (Zea mays L.), that are expressed specifically in mesophyll and bundle-sheath cells, respectively [231], where only the isoform II allowed cyclic electron transfer when introduced into the cyanobacterium Plectonema boryanum. In reverse genetics experiments the two leaf ferrodoxin isoforms from A. thaliana were analyzed [232]. Interestingly LEF is retarded in lines with lower Fd 2 (the most abundant Fd species) levels and controversially under certain conditions enhanced in lines with lower Fd 1 (the minor isoprotein) levels. These data indicate that to some degree the roles of Fd 1 and 2 are different in photosynthetic electron transfer, pointing to a differential partitioning of electrons in LEF and CEF flow. This level of regulation could also operate in C. reinhardtii where out of the six Fds encoded in the genome, five are localized in the chloroplast [233,234]. Acknowledgments A.B. acknowledges funding from the DFG (BU2536/1-1). M.H. acknowledges funding from the DFG. References [1] N. Nelson, Plant photosystem I—the most efficient nano-photochemical machine, J. Nanosci. Nanotechnol. 9 (2009) 1709–1713.

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