Journal of Plant Physiology 169 (2012) 1639–1653
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Journal of Plant Physiology
journal homepage: www.elsevier.de/jplph
Structure and function of photosystem I and its application in biomimetic
solar-to-fuel systems
Joanna Kargul ∗ , Julian David Janna Olmos, Tomasz Krupnik
Department of Plant Molecular Physiology, Faculty of Biology, University of Warsaw, ul. Miecznikowa 1, 02-096 Warsaw, Poland
a r t i c l e
i n f o
Article history:
Received 30 January 2012
Received in revised form 9 May 2012
Accepted 11 May 2012
Keywords:
Artificial photosynthesis
Hydrogen
Light harvesting complex I
Photosystem I
Photosynthetic reaction centre
a b s t r a c t
Photosystem I (PSI) is one of the most efficient biological macromolecular complexes that converts solar
energy into condensed energy of chemical bonds. Despite high structural complexity, PSI operates with
a quantum yield close to 1.0 and to date, no man-made synthetic system approached this remarkable
efficiency. This review highlights recent developments in dissecting molecular structure and function
of the prokaryotic and eukaryotic PSI. It also overviews progress in the application of this complex as a
natural photocathode for production of hydrogen within the biomimetic solar-to-fuel nanodevices.
© 2012 Elsevier GmbH. All rights reserved.
Introduction
Oxygenic photosynthesis, the fundamental process of conversion of sunlight into chemical energy, sustains life on earth. The
first step in this process, the light-driven charge separation, is
conducted by photosystems (PS) I and II, two large multimeric
chlorophyll (Chl)-binding protein complexes embedded in the thylakoid membranes of cyanobacteria, algae and higher plants. PSI
and photosystem II (PSII) evolved from a common ancestor and
are constructed around an exquisitely designed basic blueprint.
Both contain a reaction centre (RC) protein complex coupled to
a light harvesting (LH) system made up of several hundred pigment molecules (Blankenship, 2010; Kargul and Barber, 2011). The
energy of photons captured by the LH systems of PSII and PSI is
rapidly transferred to the photochemical reactions centres, the
so-called P680 and P700 Chla molecules, respectively, where it
powers the vectorial movement of electrons across a membrane,
thus generating an electrical gradient, as well as a chemical potential gradient in the form of ‘redox’ energy (Kargul and Barber, 2011).
The P680+ cation is the strongest, most abundant oxidizing species
Abbreviations: Asc, ascorbate; CET, cyclic electron transport; Chl, chlorophyll;
cyt, cytochrome; DCPIP, 2,6-dichlorophenolindophenol; ETC, electron transfer
cofactors; Fd, ferredoxin; FNR, ferredoxin-NADP+ reductase; H2 ase, hydrogenase; LHCI, light harvesting complex of PSI; LHCII, light harvesting complex of
PSII; MA, mercaptoacetic acid; MV, methyl viologen; PC, plastocyanin; PMS, Nmethylphenazonium methyl sulphate; PSI, photosystem I; PSII, photosystem II; RC,
reaction centre; ROS, Reactive Oxygen Species.
∗ Corresponding author. Tel.: +48 225542005; fax: +48 225543910.
E-mail address: [email protected] (J. Kargul).
0176-1617/$ – see front matter © 2012 Elsevier GmbH. All rights reserved.
http://dx.doi.org/10.1016/j.jplph.2012.05.018
known in biology (with a midpoint potential estimated at +1.25 eV)
(Rappaport et al., 2002; Grabolle and Dau, 2005) that generates the
most positive redox potential found in natural systems required for
the thermodynamically demanding reaction of water oxidation. In
contrast, P700 upon light excitation generates the most powerful
naturally occurring reductant in the form of P700* (Em ∼ −1.26 eV)
(Blankenship, 2002). In this way, PSI largely determines the global
amount of enthalpy achievable in the living systems (Nelson, 2011).
The photosynthetic RCs P680 and P700 are coupled, so as to use
two photons to drive each electron through the system, providing
sufficient energy to oxidize water and to reduce CO2 . Photocatalytic oxidation of substrate water molecules conducted by PSII
generates not only the reducing equivalents used for production
of biomass, but also the vast majority of molecular dioxygen that
sustains the aerobic atmosphere on our planet. The redox coupling
between both RCs occurs through the action of the cytochrome b6 f
complex (cyt b6 f), which donates water-derived electrons to a soluble electron carrier plastocyanin (PC) or cytochrome c6 (cyt c6 ),
and maintains formation of the electrochemical gradient across the
thylakoid membrane.
Photosystem I (PSI) catalyzes the light-driven vectorial electron
transfer from PC or cyt c6 at the lumenal side of the thylakoid membrane, to ferredoxin (Fd) at the stromal side. Under some stress
conditions that lead to cellular ATP depletion, a cyclic electron flow
(CEF) around PSI is activated, whereby the reduced acceptors of
PSI donate electrons to the cyt b6 f complex to produce exclusively
ATP and proton gradient across the thylakoid membrane. Although
significant progress has been made in identifying the molecular
components of CEF, the precise molecular pathways and physiological relevance of this process in various types of photosynthetic
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organisms, especially in higher plants, are still a matter of debate. As
this regulatory process is beyond the scope of this review, readers
are referred to some excellent recent reviews on this topic (Kramer
et al., 2004; Shikanai, 2007; Alric, 2010; Peltier et al., 2010; Johnson,
2011; Rochaix, 2011).
Although a wide range of techniques has been applied to obtain
structural and functional details of the PSI and PSII RCs and all
the other molecular components of the photosynthetic machinery,
by far the most informative has been X-ray crystallography. There
are now medium-to-high resolution crystal structures of prokaryotic PSI (Jordan et al., 2001), eukaryotic PSI supercomplex with
its light harvesting antenna system (LHCI-PSI) (Ben-Shem et al.,
2003; Amunts et al., 2007, 2010), prokaryotic PSII (Zouni et al.,
2001; Kamiya and Shen, 2003; Ferreira et al., 2004; Loll et al., 2005;
Guskov et al., 2009; Umena et al., 2011), cyt b6 f (Kurisu et al., 2003;
Stroebel et al., 2003; Yan et al., 2006; Yamashita et al., 2007; Baniulis
et al., 2009), trimeric and monomeric light harvesting antenna systems of PSII (LHCII) (Liu et al., 2004; Standfuss et al., 2005; Pan
et al., 2011), and the soluble electron carriers, including PC (Guss
and Freeman, 1983; Moore et al., 1991; Redinbo et al., 1993), cyt
c6 (Frazao et al., 1995; Kerfeld et al., 1995), Fd (Morales et al.,
1999; Kameda et al., 2011) and ferredoxin-NADP+ reductase (FNR)
(Karplus et al., 1991; Serre et al., 1996; Deng et al., 1999; Kurisu
et al., 2001; Tejero et al., 2003; Muraki et al., 2010). These structures
have facilitated great advancement in our detailed mechanistic
knowledge of photosynthetic electron transfer, and in particular,
the fundamental processes of photocatalytic water splitting and
charge separation in the PSI and PSII RCs (Haumann et al., 2005;
Brudvig, 2008; Cogdell et al., 2008; Sproviero et al., 2008; Siegbahn,
2011).
The LH antenna systems associated with RCs of PSI and PSII
allow them to operate efficiently under relatively low light intensities. The nature of the LH systems varies considerably among
phototrophs, but all function to intercept light and transfer the
excitation energy rapidly to the photochemical RCs. In order for
the process to be efficient, the overall transfer rate must be faster
than the singlet lifetimes of the pigments, which are typically in the
nanosecond time domain (Cogdell et al., 2008). In fact, overall transfer times of energy migration from the LH system to the RC are in
the sub-nanosecond time domain (Cogdell et al., 2008; Collins et al.,
2011). The different spectral properties of the wide range of LH pigments, coupled with fine-tuning by interactions with the proteins
to which they bind, allow photosynthetic organisms to absorb at all
the wavelengths available in the visible part of the solar spectrum
at the Earth’s surface spanning between 350 and 1000 nm (Archer
and Barber, 2004; Blankenship et al., 2011).
Structure and function of PSI
Protein subunits and bound cofactors
The two major breakthroughs in revealing the detailed molecular organization of PSI were the 2.5-Å X-ray structure of
prokaryotic PSI from the cyanobacterium Thermosynechococcus
elongatus (Jordan et al., 2001; shown in Fig. 1A) and the 4.4-Å
X-ray structure of the eukaryotic PSI complex with its associated
light-harvesting antenna system from pea (Pisum sativum) that was
subsequently refined to 3.3 Å (Ben-Shem et al., 2003; Amunts et al.,
2007, 2010; see Fig. 1B). Detailed structural comparison of the protein backbone, inbound redox cofactors and the pigment systems
in the prokaryotic and eukaryotic PSI complexes that are separated
1.5 billion years apart provided important insights into the evolution of this complex (reviewed in Schubert et al., 1998; Nelson and
Ben-Shem, 2005; Sadekar et al., 2006; Amunts and Nelson, 2009;
Nelson, 2011).
Fig. 1. X-ray crystal structures of prokaryotic and eukaryotic PSI. (A) 2.5-Å X-ray
crystal structure of the cyanobacterial PSI from T. elongatus. Shown are the helices of
several of the protein subunits, PsaA (yellow), PsaB (blue), PsaC (green), PsaD (cyan),
PsaE (orange), PsaL (red), PsaF (grey), PsaK (magenta) and the cofactors (green apart
from [4Fe–4S] clusters shown in red). For clarity, some subunits and cofactors are
omitted. Only one monomer of the biologically active trimer is shown. The view is
approximately with the threefold symmetry axis of the protein in the membrane
plane. (B) 3.3 Å X-ray crystal structure of the higher plant LHCI-PSI from P. sativum.
View is from the stromal side. Shown are the helices of several of the protein subunits
with color coding as above, as well as 3 novel core subunits: PsaG (wheat), PsaH
(cyan), and PsaN (pink) and the Lhca1-Lhca4 antenna subunits (green). For clarity,
the stromal extrinsic subunits and some small core subunits are not shown. Figure
produced from PDB coordinates 1JB0 (Jordan et al., 2001) and 3LW5 (Amunts et al.,
2010) using the MBT Protein Workshop software (Moreland et al., 2005).
Cyanobacterial PSI exists predominantly as trimers in vivo
(Kruip et al., 1994; Karapetyan et al., 1999; Jordan et al., 2001),
although monomeric, dimeric and tetrameric forms of this complex have also been reported in prokaryotic phototrophs (Rögner
et al., 1990; Kruip et al., 1994; Karapetyan et al., 1999; El-Mohsnawy
et al., 2010; Watanabe et al., 2011). In contrast, eukaryotic PSI is
always monomeric (Scheller et al., 2001; Busch and Hippler, 2011).
The LHCI-PSI supercomplex has a molecular mass of ∼600–770 kDa
(including the inbound cofactors) and is composed of two structural
and functional domains: the core (or reaction centre complex of
estimated molecular mass of 310–356 kDa excluding the inbound
cofactors), in which the bulk of light capturing and charge separation occurs, and the external (peripheral) antenna system that
increases the light-harvesting capacity of PSI and transmits quanta
of solar energy to the core antenna, and ultimately, to the P700
RC. While the PSI core is highly conserved throughout evolution, with differences mainly in some small intrinsic core subunits
(Jordan et al., 2001; Ben-Shem et al., 2003), the light-harvesting
system varies considerably between species with respect to its
subunit and pigment composition, and stoichiometry, reflecting
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
evolutionary adaptation of photosynthetic organisms to diverse
ecological niches (Collins et al., 2011; Busch and Hippler, 2011).
In green algae and higher plants, the outer antenna is composed of at least 4 nuclear-encoded Chla/b-binding Lhca proteins
that form the crescent-shaped light harvesting complex I (LHCI)
asymmetrically attached to the core domain (Scheller et al., 2001;
Ben-Shem et al., 2003; Kargul et al., 2003, 2005; Drop et al., 2011;
Busch and Hippler, 2011). In chromalveolates, the LHCI complex
is formed by the Chla/c-binding proteins, for example the diatom
fucoxanthin-chlorophyll-proteins (FCPs), which bind Chlc instead
of Chlb and fucoxanthin instead of lutein (Durnford et al., 1999;
Veith and Büchel, 2007; Veith et al., 2009; Neilson and Durnford,
2010). In the red algae Rhodophyta the LHCI antenna is composed
of a varying number of exclusively Chla-binding Lhcr proteins
(Gardian et al., 2007; Busch et al., 2010; Thangaraj et al., 2011; see
Fig. 2A). For a more detailed overview of the various types of the
LHCI antenna systems, the reader is advised to refer to an excellent
recent review by Busch and Hippler (2011).
In the crystallographic model of a higher plant LHCI-PSI supercomplex, the outer LHCI antenna “belt” is composed of 4 Lhca
subunits that asymmetrically attach to the core domain on the
PsaF/PsaJ side, as shown in Fig. 1B (Ben-Shem et al., 2003; Amunts
et al., 2007, 2010). This is the central building block of the plant
and green algal outer light-harvesting antenna, which is organized
as two functional heterodimers composed of Lhca1–Lhca4 and
Lhca2–Lhca3 subunits (Croce et al., 2002; Ben-Shem et al., 2003),
although 2–7 additional Lhca subunits may associate with this basic
peripheral antenna system (Germano et al., 2002; Kargul et al.,
2003, 2005; Ganeteg et al., 2004; Storf et al., 2004; Lucinski et al.,
2006; Stauber et al., 2009; Drop et al., 2011).
In addition to its function in light harvesting, the LHCI complex has an important role in photoprotection, since the production
of the Reactive Oxygen Species (ROS) and photoinhibition are
unavoidable outcomes of oxygenic photosynthesis at all light intensities (Ruban and Johnson, 2010). LHCI has been suggested to act
as a safety valve under the conditions of high light illumination,
when overexcitation of both photosystems can occur. This is due to
the presence of the red-shifted Chl molecules in the LHCI complex
and the PSI core antenna that absorb light above 700 nm and act
as excitation sinks enabling an uphill energy transfer to the P700
reaction centre (Croce et al., 2000; Ihalainen et al., 2002; Jennings
et al., 2003; Melkozernov et al., 2004, 2005).
In cyanobacteria, the peripheral light harvesting antenna is
formed by large water-soluble phycobilisome complexes that
attach to the PSI core on the stromal side of the thylakoid membrane (Glazer, 1985; MacColl, 2004; Murray et al., 2006). Under
some conditions such as low availability of iron, the peripheral
antenna system is dominated by an intrinsic Chla-binding protein
complex made up of 1–2 rings of 18–43 copies of the CP43-like
IsiA subunits (Bibby et al., 2001a,b; Boekema et al., 2001; Nield
et al., 2003; Yeremenko et al., 2004; Kouril et al., 2005a; Chauhan
et al., 2011) that enclose trimeric PSI. A similar antenna system is
found in the cyanobacteria known as prochlorophytes, which use
Chla/Chlb-binding Pcb proteins, as shown in Fig. 2D (Bibby et al.,
2001c, 2003a,b). The Pcb and IsiA proteins are structurally related
to each other and to CP43, CP47 and the N-terminal domains of the
PSI RC proteins, PsaA and PsaB (La Roche et al., 1996; Green and
Durnfold, 1996; Barber et al., 2006).
Iron limitation leads to fast degradation of photosynthetic RCs,
which all contain iron in haem and [4Fe–4S] clusters. The most
severely affected is cyanobacterial PSI, as it is the largest sink of
iron, with 36 Fe atoms in nine [4Fe–4S] clusters per PSI trimer,
associated with additional [2Fe–2S] clusters in multiple copies of
Fd (Jordan et al., 2001). During iron deprivation, induction of IsiA
expression occurs by de-repressing the isiAB operon which encodes
the IsiA (CP43 ) antenna subunit and IsiB (flavodoxin) proteins.
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The latter functionally replaces Fe-containing Fd. The main role
of the multiple IsiA rings is to significantly increase the functional
antenna or absorption cross-section, which compensates for degradation of PSI RCs under Fe limitation (Andrizhiyevskaya et al., 2002;
Melkozernov et al., 2003; Chauhan et al., 2011).
Significant remodeling of the photosynthetic apparatus upon
iron starvation also occurs in photosynthetic eukaryotes. Iron limitation in the green alga Chlamydomonas reinhardtii results not
only in pronounced degradation of the PSI RCs, but also in gradual dissociation of the LHCI antenna subunits and a transient
increase of the functional LHCII antenna, the latter acting as a pigment buffer and storage (Moseley et al., 2002; Naumann et al.,
2005, 2007). A similar phenomenon is observed in red algae and
diatoms, whereby the iron stress induces reduction of the functional antenna of PSI (Desquilbet et al., 2003; Doan et al., 2003; Allen
et al., 2008; Juhas and Büchel, 2012). Thus, a decrease of the PSI
absorption cross-section seems to serve as a general mechanism of
reducing photo-oxidative damage to the eukaryotic photosynthetic
apparatus.
While the cyanobacterial PSI core complex in the absence of
IsiA or Pcb proteins comprises 12 subunits harboring 96 Chls per
monomer (Jordan et al., 2001), the higher plant PSI is considerably larger, containing at least 15 core subunits (PsaA to PsaL and
PsaN to PsaP, with the PsaO and PsaP proteins absent in the currently available the X-ray structures), at least 4 stably associated
Lhca antenna subunits and a total of 173 Chl and 15 carotenoid
molecules, as shown in Fig. 1B (Ben-Shem et al., 2003; Amunts et al.,
2007, 2010). Most of the additional Chls present in the plant LHCIPSI supercomplex over and above those within the cyanobacterial
PSI are associated with the four Lhca antenna subunits or belong to
the “linker” and “gap” Chls that facilitate energy transfer within the
LHCI antenna and between the LHCI antenna and the RC (Ben-Shem
et al., 2003). The twelve subunits of the cyanobacterial core domain
include 9 intrinsic polytopic subunits (PsaA, PsaB, PsaF, PsaI, PsaJ,
PsaK, PsaL, PsaM and PsaX) and 3 extrinsic stromal subunits (PsaC,
PsaD and PsaE), as presented in Fig. 1A.
The central part of the core complex is formed by a highly conserved heterodimer of the PsaA and PsaB subunits which bind the
majority of the electron transfer cofactors (ETCs), antenna and lipid
cofactors. The N-termini of both RC subunits are oriented toward
the stroma, whereas the C-termini are exposed toward the thylakoid lumen. Both subunits contain 11 transmembrane helices
that are divided into the N-terminal domain composed of six ␣helices (A/B-a to A/B-f) and a C-terminal domain containing five
␣-helices (A/B-g to A/B-k; nomenclature according to Jordan et al.,
2001). The latter form two interlocked semicircles enclosing the
ETCs, including 6 Chla molecules, two phylloquinones, and a single [4Fe–4S] iron–sulphur cluster, termed FX (Jordan et al., 2001;
Ben-Shem et al., 2003). Two other [4Fe–4S] clusters (FA and FB ) are
bound to the PsaC subunit located on the stromal side of the complex. This subunit is evolutionarily related to the class of bacterial
Fds (Antonkine et al., 2003). Fifteen ␤-carotenoids were identified in the 3.3 Å structure of the higher plant PSI (Amunts et al.,
2010), whereas 30 ␤-carotenoids were built into the model of the
cyanobacterial PSI (Jordan et al., 2001). All the ETCs are arranged
in two symmetric branches along the crystallographic pseudo-C2
axis, as depicted in Fig. 3. The other intrinsic subunits are peripheral to the PsaA/B heterodimer and coordinate some of the inner
antenna cofactors (Jordan et al., 2001; Amunts et al., 2010).
While the higher plant core domain retains the location and orientation of the ETCs and its protein backbone is structurally very
similar to the cyanobacterial PSI RC, it does not have the X and M
subunits. Instead, 4 additional core subunits are present exclusively
in higher plants and green algae, namely subunits PsaG, PsaH, PsaN,
and PsaO (Scheller et al., 2001; Green and Durnfold, 1996; Knoetzel
et al., 2002). These subunits play specific roles in association of the
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Fig. 2. Top view projections of various types of LHC-PSI supercomplexes derived from electron microscopy and single particle analysis. (A) Stromal top view of the LHCI-PSI
supercomplex from the thermoacidophilic red alga Galdieria sulphuraria with an overlay of the X-ray structure of plant LHCI-PSI (Thangaraj et al., 2011). The crescent-shaped
area highlighted in green is suggested to contain 4–5 additional Lhcr subunits over and above the four Lhca subunits in the plant X-ray structure. Scale bar, 10 nm. (B)
Projection map for the LHCI–PSI supercomplex isolated from Chlamydomonas reinhardtii cells induced to State 2 with an overlay of the plant PSI X-ray structure (Kargul
et al., 2005). Modeling is based on higher plant PDB coordinates 1QZV (Ben-Shem et al., 2003) with PSI core (green), LHCI antenna (red), PsaJ (yellow), PsaK (magenta),
PsaG (purple), PsaI (orange), PsaL (cyan) and PsaH (white). Chlorophylls are shown in yellow. The additional density observed in State 2 LHCI–PSI supercomplex is able to
accommodate an additional LHC subunit (blue) attributed to phospho-CP29. Scale bar, 5 nm. (C) Stromal top view projection map of the IsiA–PSI supercomplex isolated from
iron-starved cyanobacterium Synechocystis PCC 6803 with an overlay of the X-ray crystal structure of the trimeric PSI from T. elongatus. The supercomplex is formed by a
PSI trimer within the centre of an 18-mer ring of the Chla-containing the IsiA subunits (Bibby et al., 2001a). Scale bar, 5 nm. (D) Projection of the top view of the Pcb–PSI
supercomplex isolated from marine oxyphotobacteria Prochlorococcus marinus, strain SS120. The antenna ring of 18 Pcb Ca/b subunits surrounds a trimeric PSI reaction
centre (Bibby et al., 2001c). Scale bar, 5 nm.
LHCI antenna complex (such as PsaG; Ben-Shem et al., 2003) or
docking of a mobile fraction of the LHCII complex, during photosynthetic state transitions (subunits PsaH, and PsaO; Kargul and Barber,
2008). Interestingly, a single transmembrane helix adjacent to PsaL
and corresponding to the PsaH subunit has been shown to bind one
Chl molecule (Ben-Shem et al., 2003; see Fig. 1B). This subunit most
likely forms a docking site for the mobile LHCII complex that transiently attaches with the PSI core during state transitions, under
conditions favoring excitation of PSII (Lunde et al., 2000; Kouril
et al., 2005b; Kargul et al., 2005; Kargul and Barber, 2008). On the
opposite side of the core complex the 2-TM PsaG subunit provides
the contact surface area for the association of the belt-shaped LHCI
(Ben-Shem et al., 2003), as shown in Fig. 1B.
The precise role of the lumenal PsaN subunit is still debatable. In
the latest 3.3-Å X-ray structure of plant LHCI-PSI, it seems to interact with the Lhca2 and Lhca3 subunits, thus potentially stabilizing
the formation of this antenna heterodimer (Amunts et al., 2010). It
has also been suggested to be indirectly involved in binding of PC
by providing the correct orientation of the N-terminal domain of
PsaF, the docking site for this electron carrier (Haldrup et al., 1999,
2000; Jensen et al., 2007), although the 3.3-Å X-ray structure of
LHCI-PSI seems to preclude the direct interaction of PsaN and PsaF.
PsaN has also been proposed to bind the minor Lhca subunit Lhca5
(Storf et al., 2005; Lucinski et al., 2006) which, together with Lhca6,
is important for the formation of the PSI-NADH dehydrogenase-like
(NDH) complex during the cyclic electron transport (CET) around
PSI in higher plants (Peng et al., 2008, 2009; Peng and Shikanai,
2011).
Interestingly, some cyanobacterial core subunits have been
retained in the PSI core of the eukaryotic chromalveolates and
red algae. In contrast to higher plants, the plastid genome of
the diatom P. tricornutum contains the PsaM subunit, which is
suggested to enable trimerization of PSI in cyanobacteria and facilitate energy transfer between monomers of the cyanobacterial
PSI trimer (Grotjohann and Fromme, 2005), albeit with sequence
identity of only 50% (Veith and Büchel, 2007). Similar to diatoms,
red algae retained the cyanobacterial PsaM subunit, as shown by
the recent plastid genome analysis of a primitive unicellular red
microalga Galdieria sulphuraria (Vanselow et al., 2009).
Notably, some higher plant PSI RC subunits are absent in other
eukaryotic PSI complexes. The PsaG subunit that provides a docking site for the LHCI antenna assembly in higher plants and green
algae has not been identified in diatoms (Armbrust et al., 2004) or
red algae (Matsuzaki et al., 2004; Vanselow et al., 2009). Similarly,
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
Fig. 3. Electron transfer cofactors of PSI. Arrangement of the electron transport
cofactors (ETCs) in cyanobacterial PSI from T. elongatus viewed along the membrane
plane. The cofactors of the ETCs are related by the pseudo-symmetry C2 axis passing through FX , and oriented normal to the paper plane. Nomenclature according to
Jordan et al. (2001). eC-A1/B1: a primary electron donor; eC-A2/B2, accessory Chls;
eC-A3/B3 Chls, A0 primary electron acceptor in P700; QK -A and QK -B, A1 phylloquinones which are secondary electron acceptors in P700; FX , FA , FB : [4Fe–4S] clusters,
the latter two shown within the backbone of PsaC (pink). Figure produced from PDB
coordinates 1JB0 (Jordan et al., 2001) using the PyMOL molecular graphics system
(DeLano, 2002).
the PsaH subunit, important in state transitions in green algae and
higher plants, is also absent in the genomes of diatoms and red
algae. In addition, green algae lack a gene coding for PsaP (Jensen
et al., 2007).
In red algae, the PsaL and PsaF core subunits are of a chimeric
nature. While the cyanobacterial PsaL, with its extended Cterminus protruding from the RC, is essential for PSI trimer
formation (Chitnis and Chitnis, 1993; Jordan et al., 2001), its higher
plant counterpart forms, together with the PsaH subunit, the
mobile LHCII docking site during the adaptation process of state
transitions (Haldrup et al., 2000). This extended C-terminal domain
of PsaL is absent in plants and algae (Ben-Shem et al., 2003), in
agreement with the exclusively monomeric character of eukaryotic PSI. The conserved sequences required for PSI trimerization in
cyanobacteria (the C-terminal helix and the Ca2+ -binding site) and
the plant-specific motifs for PsaH interaction are absent in the PsaL
subunit of G. sulphuraria (Vanselow et al., 2009).
The chimeric nature of PsaF in G. sulphuraria is emphasized by
a striking homology of its N-terminal domain to the higher plant
counterpart and the presence of the conserved cyanobacterial-like
motifs at the C-terminus. The N-terminal domain of G. sulphuraria
PsaF contains the positively charged Lys-rich motif that is essential for tight docking of PC and cyt c6 in plants and green algae
(Hippler et al., 1996, 1998, 1999). The conserved C-terminal extension of G. sulphuraria PsaF has been suggested to interact with the
peripheral antenna composed of both LHCI subunits and phycobilisomes (Vanselow et al., 2009), although the latter remains to
be confirmed experimentally. Similar to cyanobacteria, G. sulphuraria and other red algae such as Cyanidioschyzon merolae lack a
gene for PC, but exclusively use the more ancient cyt c553 as a soluble electron carrier donating electrons to the photo-oxidized P700+
(Matsuzaki et al., 2004; Vanselow et al., 2009).
The majority of the core pigments (87 and 90 Chla, as well as 15
and 22 ␤-carotenoid molecules in higher plants and cyanobacteria,
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Fig. 4. Structural similarity of the N-terminal domain of PSI and PSII RC. The
N-terminal domain of the PSI RC heterodimer composed of 6 TMs of the inner peripheral antenna of PSI is aligned with the CP43/CP47 inner antenna complex of PSII.
Shown are the backbones of the protein subunits: D1 (cyan), D2 (blue), CP43 (red),
CP47 (orange), N-terminal domain of PsaA (deep purple) and N-terminal domain of
PsaB (yellow). The views are approximately with the twofold symmetry axis of the
protein in the membrane plane. The coordinates are 1S5L (Ferreira et al., 2004), and
1JB0 (Jordan et al., 2001). Figure produced from PDB coordinates using the PyMOL
molecular graphics system (DeLano, 2002). Figure adapted from Kargul and Barber
(2011).
respectively) act as light-harvesting antenna coordinated mainly
by the PsaA/B heterodimer aided by some small intrinsic core subunits (Jordan et al., 2001; Ben-Shem et al., 2003; Amunts et al.,
2007, 2010). Two types of inner antenna can be distinguished in the
PSI RC: the innermost ‘central antenna’ of 43 Chls forming a circle
around the ETCs at a distance not less than ∼18 Å flanked by 2 layers
of ‘peripheral inner antenna’ of 18 Chls each that are bound predominantly to the N-terminal domains of the PsaA/B heterodimer
(Jordan et al., 2001). The crystal structure of plant LHCI-PSI indicates
that the majority of the RC Chls retained the same position and tilting angle as in the cyanobacterial PSI, indicating a high degree of
conservation of the PSI RC even after 1.5 billion years of evolution
(Jordan et al., 2001; Ben-Shem et al., 2003).
The structures of the N-terminal domains of PsaA and PsaB are
equivalent to those of CP43 and CP47 inner antenna subunits of PSII,
whereas, the organization of the five TM helices of the C-terminal
domains of PsaA and PsaB is similar to that of the D1 and D2 subunits of the PSII RC (Schubert et al., 1998; Barber et al., 1999; Kargul
and Barber, 2011; see Fig. 4), indicating a common evolutionary
origin of both types of RCs (Ben-Shem et al., 2004; Nelson and
Ben-Shem, 2005; Amunts and Nelson, 2009; Hohmann-Marriott
and Blankenship, 2011; Kargul and Barber, 2011). Moreover, distribution of the peripheral antenna Chls bound to the N-terminal
domains of PsaA and PsaB is similar to that of the CP43 and CP47.
The C-terminal domains of PsaA and PsaB jointly coordinate 25 Chla
molecules of the peripheral and central antenna (Jordan et al., 2001)
whose position is optimized to mediate energy transfer to P700 and
fast trapping of excitation energy in the PSI RC.
On the lumenal (donor) side of PSI, the docking site for the
mobile electron donors, cyt c6 or PC is formed by the ␣-helices of
loops A/B-ij of the PsaA/B heterodimer containing 2 conserved Trp
residues, as shown in Fig. 5. The plant PsaF subunit contains the
N-terminal domain that is longer compared to its cyanobacterial
counterpart. This extended domain forms the amphipathic Lys-rich
‘helix–loop–helix’ motif (Ben-Shem et al., 2003) that enables strong
electrostatic interaction with the acidic regions (negative patch) of
PC and, as a consequence, two orders of magnitude faster electron
transfer from this mobile electron carrier to P700+ in higher plants
compared to cyanobacteria (Hippler et al., 1996, 1998).
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J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
transfer between the [2Fe–2S] cluster of Fd and the distal FB cluster of PSI (Fischer et al., 1998, 1999; Ruffle et al., 2000; Bottin et al.,
2001; Sétif et al., 2002; Antonkine et al., 2003).
The P700 RC and electron transfer chain
Fig. 5. Structural basis of PSI interaction with plastocyanin and ferredoxin. Presented is 3.3-Å X-ray crystal structure of higher plant photosystem I (Amunts et al.,
2010; PDB coordinates 3LW5). Shown are P700 reaction centre and the electron
transfer cofactors (color coding as in Fig. 3) coordinated by subunits PsaA (yellow)
and PsaB (blue). For clarity all but two of 11-transmembrane helices of the PsaA and
PsaB protein bundle are omitted. At the vertex of the P700-coordinating helices
shown are Trp658 of PsaA and Trp625 of PsaB (both in black) forming lumenal
hydrophobic patch. Functionally important N-terminal Lys residues (Lys16, Lys23
and Lys30; Hippler et al., 1998) of PsaF (gray) are indicated. They form a positively
charged patch of electrostatic potential (marked as blue mesh). Both electrostatic
and hydrophobic interactions are important for docking of cyt c6 or plastocyanin,
PC (cyan). Plastocyanin 1.50-Å X-ray crystal structure (Redinbo et al., 1993; PDB
coordinates 2PLT) shows two negatively charged patches Asp42/Glu43/Asp44 and
Asp59/Asp60, generating negatively charged electrostatic potential (red mesh).
Mutation in the first patch impairs binding to PSI (Hippler et al., 1996). Binding of
electron donor occurs when the negatively charged residues in the first patch interact with positively charged residues in the PsaF subunit aided by the hydrophobic
area formed by Trp659 and Trp625 residues of the PsaA/B heterodimer. The electrons
are then energized and transported further to iron–sulphur clusters in sequence
FX , FA , and FB (red cubes) where they arrive at the electron acceptor docking side
formed by extrinsic subunits PsaC, PsaD and PsaE. All three units contribute to the
ferredoxin (Fd) docking with positively charged residues. The key-charge residues
(black) in PsaC are Lys35/Gly36/Lys51/Arg52, in PsaC His97/Asp98/Lys103/Arg108
and PsaE Arg40. Those residues form a dent on the stromal side of PSI and generate positively charged electrostatic potential shown as blue mesh. The 1.46-Å
X-ray crystal structure (Kameda et al., 2011, PDB coordinates 3AV8) of Fd (magenta)
is placed in the vicinity of the docking site and faces toward PSI with negatively
charged residues generating electrostatic potential (shown as red mesh). Mutations
in residues Asp66/Asp67 (not shown) deplete efficiency of binding with FNR. The
iron–sulphur cluster of Fd acts as the electron acceptor centre (red rectangle).
On the stromal (acceptor) side of PSI, the binding pocket for
the mobile electron acceptors Fd or flavodoxin is formed jointly
by the extrinsic PsaC, PsaD and PsaE subunits, as shown in Fig. 5.
These three stromal subunits, which are highly conserved throughout evolution, play distinctive roles in binding of Fd, with PsaD
providing the electrostatic guidance of Fd into the PSI binding
pocket, PsaE (with its Arg39) stabilizing the molecular electron
transfer complex of Fd and PSI, and PsaC (with its Lys35), forming
a close protein-protein interaction that is essential for fast electron
As briefly discussed in the previous section, comparison of the
crystallographic structures of cyanobacterial and higher plant PSI
complexes shows a remarkable similarity in the organization and
specific binding sites of the ETCs. This functionally most important
part of PSI is formed by six Chla molecules, two phylloquinones
and three [4Fe–4S] clusters, as shown in Fig. 3. The Chls and phylloquinones are arranged along two branches, A and B, as pairs of
pseudo-dimers related by the pseudo-symmetry C2 axis and coordinated to the side chains of the PsaA/B heterodimer (Jordan et al.,
2001; Ben-Shem et al., 2003). Branch A is composed of Chls eC-A1,
eC-B2, eC-A3 and a phylloquinone QK -A, whereas branch B contains
Chls eC-B1, eC-A2, eC-B3 and a phylloquinone QK -B (nomenclature
according to Jordan et al., 2001). The two branches join again at
the [4Fe–4S] cluster FX which is followed by the two additional
[4Fe–4S] clusters FA and FB , both of which are coordinated by side
chains of the stromal extrinsic subunit, PsaC.
It is now widely accepted that both branches in the PSI RC are
active in electron transfer, albeit operating with different kinetics,
as shown by numerous transient optical spectroscopy measurements coupled to mutagenesis of the ETC coordinating ligands
(Guergova-Kuras et al., 2001; Ramesh et al., 2004; Bautista et al.,
2005; Dashdorj et al., 2005; Poluektov et al., 2005; Santabarbara
et al., 2005, 2008; Li et al., 2006; Müller et al., 2010). The rate
constants are 35 × 106 × s−1 and 4.4 × 106 s−1 for the electron transfer steps from each phylloquinone to FX (Joliot and Joliot, 1999;
Guergova-Kuras et al., 2001). It appears that branch A is about
10-fold slower than branch B, despite the faster initial charge separation (Guergova-Kuras et al., 2001; Müller et al., 2010). Although
both branches participate in electron transfer, branch A appears to
be a dominating one with a branch A/branch B ratio varying from
3:3 in green algae to ∼4:1 in cyanobacteria (Ramesh et al., 2004;
Dashdorj et al., 2005; Li et al., 2006; Müller et al., 2010).
The photochemical RC of PSI comprises a cluster of 6 Chla
molecules that function as the primary electron donors and primary electron acceptors. The primary electron donor of PSI RC, the
so-called P700 (Em of ∼0.5 eV), is formed by the ‘special’ pair of
the eC-A1/eC-B1 Chls that are excitonically tightly coupled with a
Mg–Mg distance of 6.6 Å (Jordan et al., 2001). The chlorin planes of
the P700 Chls are oriented perpendicular to the membrane plane
and form a stacked dimer with a 3.6 Å interplanar distance. This
organization varies from the ‘special pair’ of the purple bacterial
RC where the Mg–Mg distance is larger at 7.6 Å (Allen et al., 1987;
Deisenhofer et al., 1995). In contrast to the homodimeric bacterial
special pair, P700 Chls form a heterodimer, with eC-A1 being Chla
13 -epimer (Watanabe et al., 1985; Jordan et al., 2001). The heterodimeric nature of P700 primary electron donor is also reflected
by the presence of hydrogen bonds within the binding pocket of
eC-A1 and lack of those in the binding site of eC-B1 (Jordan et al.,
2001).
The other two Chla pairs are composed of the eC-B2/eC-A2
and eC-A3/eC-B3 (Jordan et al., 2001). The eC-A/B-2 Chls represent the so-called ‘accessory’ Chls, which despite being resolved
in the X-ray structures were functionally resolved by spectroscopic methods only recently (Slavov et al., 2008; Müller et al.,
2010). The eC-A/B-3 Chls are commonly referred to as A0 (Em of
−1.0 eV), which represents the primary electron acceptor reduced
in less than 10 ps, as observed by the spectroscopic measurements
(Santabarbara et al., 2010). Recently, Holzwarth and colleagues proposed an alternative mechanism for the primary charge separation,
whereby the first radical pair would form within the accessory Chls
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
1645
Table 1
Rates of hydrogen production from various types of PSI-hybrid nanodevices.
PSI-biomimetic nanoconstruct
Maximum rate of H2 production
(␮mol H2 mg Chl−1 h−1 )
References
Covalently linked PC to platinized S. oleracea PSI
PSI and T. roseopersicin [NiFe]-H2 ase in-solution system
PSI-R. eutropha [NiFe]-H2 ase hybrid complex
Platinized S. oleracea PSI
Au nanoparticle wired to PsaC-rebuilt Synechococcus sp. PCC 7002 PSI
Platinized T. elongatus PSI, cyt c6
In vitro PC/PSI/PetF/HydA1 reconstitution system
Pt nanoparticle wired to PsaC-rebuilt PSI of Synechococcus sp. PCC 7002, cyt c6
PC-PSI-Pt nanoparticle complex with 1,6-hexanedithiol molecular wire
Cobaloximized S. leoploliensis/S. lividus PSI, cyt c6
PC-PSI-Pt nanoparticle complex with 1,4-benzenedithiol molecular wire
cyt c6 -PSI-C. acetobutylicum [FeFe]-H2 ase complex
Au electrode-immobilized PSI-R. eutropha [NiFe]-hydrogenase hybrid complex
0.09
0.50
0.58
2.0
3.4
5.5
16
49.3
100.6
246
312
2200
3000
Evans et al. (2004)
Qian et al. (2008)
Ihara et al. (2006)
Millsaps et al. (2001)
Grimme et al. (2008)
Iwuchukwu et al. (2010)
Winkler et al. (2009)
Grimme et al. (2008)
Grimme et al. (2009)
Utschig et al. (2011a)
Grimme et al. (2009)
Lubner et al. (2011)
Krassen et al. (2009)
eC2+ eC3− of either branch rather than in the P700 ‘special pair’
(Holzwarth et al., 2006). This radical pair would then be re-reduced
by the P700. This postulate is based on ultrafast transient absorption measurements in intact LHCI-PSI particles from a green alga C.
reinhardtii (Holzwarth et al., 2006) and a higher plant Arabidopsis
thaliana (Slavov et al., 2008).
The A0 Chls are adjacent to a pair of phylloquinone molecules
(often referred to as A1 ) termed QK -A and QK -B according to
Jordan et al. (2001). The A1 phylloquinones (Em of −0.8 eV) act as
secondary electron acceptors that are rapidly reduced to the phyllosemiquinone radicals in ∼20–40 ps (Santabarbara et al., 2010). All
the amino acid ligands coordinating the eC-2 and eC-3 Chla pairs
are highly conserved within PsaA and PsaB from cyanobacteria to
higher plants, indicating that throughout evolution these interactions are essential for fine-tuning the redox potentials of the ETC.
An interesting exception is the marine cyanobacterium Acaryochloris marina which carries out oxygenic photosynthesis but contains
over 95% red-shifted Chld (with the in vivo absorbance maximum
of ∼710 nm) and only trace amounts of Chla in both photosystems
(Miyashita et al., 1996; Hu et al., 1998; Tomo et al., 2007). Chld
provides a potential selective advantage because it enables Acaryochloris to use infrared light (700–750 nm) that is not absorbed by
Chla in the far-red light-enriched habitat of this organism (Chen
and Blankenship, 2011). Consequently, the primary donor of PSI,
the so-called P740, is a dimer of Chld molecules. Similarly, Chld
replaces Chla for A0 and A1 .
The electron acceptor from A1 is the [4Fe–4S] cluster, the socalled FX (Evans and Cammack, 1975; Evans et al., 1976; Golbeck
et al., 1987; McDermott et al., 1989), which, similar to P700, is
located at the interface of the PsaA/B heterodimer. The FX cluster (Em of −0.7 eV) is ligated by 4 strictly conserved Cys residues
present in the loop segments A/B-hi of the PsaA/B heterodimer
(Jordan et al., 2001). The two terminal [4Fe–4S] iron–sulphur
clusters FA and FB , which operate in series, are coordinated by
Cys residues present within the conserved regions of the stromal
extrinsic PsaC subunit (Jordan et al., 2001).
PSI in biomimetic solar-to-fuel nanodevices
In-solution PSI-hybrid systems for solar-to-hydrogen production
PSI operates with a quantum yield close to 1.0 and to date,
no man-made synthetic system has approached this remarkable
efficiency. Despite high structural complexity, PSI operates as an
almost perfect Einstein photoelectric device (Nelson, 2009; Amunts
and Nelson, 2009). This means that each quantum of light harvested by the PSI antenna system reaches the P700 photochemical
RC, ultimately creating the primary radical pair species and an
ultrafast charge separation within a few picoseconds (Diner and
Rappaport, 2002; Santabarbara et al., 2010). As a result, for each
quantum of light absorbed by P700, a single electron is ejected
from the primary electron donor. These inherent light-harvesting
and electron-transfer properties of natural PSI make this macromolecular complex amenable for hydrogen production in the
solar-to-fuel biomimetic devices. As PSI forms an exceptionally
long-lived charge-separated state P700+ FB − (∼60 ms) and is characterized by an exceptionally low redox potential associated with
the distal FB cluster (Em of −0.58 eV), it provides a sufficient driving
force to reduce protons to H2 at neutral pH (Lubner et al., 2010a).
For these reasons, there is significant interest in utilizing the highly
stable natural PSI for generation of solar fuels (Blankenship et al.,
2011).
Over the last decade, the reported H2 -evolving PSI-hybrid systems consisted of solubilized individual components of electron
transfer. Consequently, the rates of electron transfer between
the interacting constituents were severely limited by diffusion.
Nonetheless, these in vitro systems have provided the chemical and
biochemical blueprints for development of the more efficient solidstate systems that operate at significantly improved H2 -evolution
rates, as summarized in Table 1 and Fig. 6.
One of the early examples of such in vitro H2 -evolving systems was based on platinization of the native PSI RCs isolated from
Spinacia oleracea (Millsaps et al., 2001). At neutral pH and room
temperature metallic platinum can be photoprecipitated on the
reducing side of PSI according to the chemical reaction:
[PtCl6 ]2− + 4e− + 4hv
Pt(s) + 6Cl−
The Hill reaction of photosynthesis, which reduces ferric ions to
ferrous ions, forms the basis for the photoprecipitation of metallic platinum onto the external surfaces of isolated PSI complexes
(Lubner et al., 2010a). Subsequently, in the four-step reducing process, PSI-derived electrons interact with Pt clusters to generate
molecular H2 .
Following illumination of the mixture containing hexachloroplatinate ([PtCl6 ]2− ), purified spinach PSI cores, PC (as a soluble
natural electron donor to photooxidized P700+ ) and an ascorbate (Asc) (as a sacrificial electron donor to PC), the Pt cations
underwent a four-step reduction to metallic Pt(s). Overall, such
a system achieved a rather modest rate of 2 ␮mol H2 mg Chl−1
h−1 (Millsaps et al., 2001). Nevertheless, it exceeded 10-fold the
efficiency of the previously reported platinized thylakoid-based
systems (Greenbaum, 1985; Lee et al., 1994, 1998).
A rather elegant diffusion-based system for biomimetic
H2 production was reported by Bruce and colleagues who
employed platinized highly stable trimeric PSI complexes from the
thermophilic cyanobacterium T. elongatus as a photocathodic component of the system (Iwuchukwu et al., 2010). By employing cyt
c6 as a natural electron donor to photooxidized PSI RC, the authors
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J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
Fig. 6. Biomimetic H2 -producing PSI nanodevices. (A) Photocatalytic system of H2 production from a PSI-cobaloxime hybrid complex. Two successive photogenerated
electrons are necessary for the catalyst to produce one H2 molecule. The electron donors depicted are Asc and cyt c6 (Utschig et al., 2011a). (B) In-solution hydrogen evolution
system composed of PSI, H2 ase, MA as electron donor and MV as electron acceptor (Qian et al., 2008). (C) Photocatalytic H2 -evolving system of platinized PSI with covalently
linked plastocyanin (PC). The covalent linker is 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (blue stick). Ascorbate serves as a sacrificial electron donor
(Evans et al., 2004). (D) PSI-molecular wire-[FeFe]-H2 ase nanodevice. The wire here is 1,6-hexanedithiol (red line). Cyt c6 was cross-linked with a zero-length cross-linking
agent to limit diffusion-based electron transfer to P700+ . The arrow indicates the directionality of electron transfer, including reduction of protons to H2 by H2 ase (Lubner
et al., 2011). (E) Bioconjugate consisting of PC cross-linked to PSI, 1,4-benzendithiol as the molecular wire (red line) and a platinum nanoparticle catalyst, DCPIP was
employed as the sacrificial electron donor (Grimme et al., 2009). (F) Au surface-immobilized PSI-[NiFe]-H2 ase photocathode. The Ni-NTA functionalized Au electrode
strongly binds His-tagged PSI (His6 -tag engineered on PsaF illustrated as a yellow anchor) and provides electrons to reduce the oxidized form of PMS as the electron donor to
P700+ . Functionality of PsaE-null mutant of Synechocystis PSI is restored by binding the MBH-PsaE fusion protein from Ralstonia eutropha. The electrons transfer from the FB
cluster at the PSI acceptor side to the distal iron–sulphur cluster of the [NiFe]-H2 ase and further to its active site, where protons are reduced to molecular hydrogen (Krassen
et al., 2009). Biomimetic devices depicted in panels A, B, D and F include the 2.5-Å X-ray crystal structure of the cyanobacterial PSI from T. elongatus (Jordan et al., 2001).
Shown are the helices of PsaA (yellow), PsaB (blue), PsaC (green), PsaD (cyan), PsaE (light brown), PsaF (grey), PsaL (red), PsaK (magenta) and the cofactors (Chls, green;
[4Fe–4S] clusters, red). Only one monomer of the biologically active trimer is shown. Panels C and E show the 3.3 Å X-ray crystal structure of higher plant PSI from P. sativum
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
reported increased efficiency of hydrogen production compared to
the system without cyt c6 . The hydrogen production, although of
a modest rate of ∼5.5 ␮mol H2 h−1 mg Chl−1 , was remarkably stable and sustained for over 85 days at temperatures elevated to
55 ◦ C, with no apparent decrease in hydrogen yield when tested
intermittently.
Recently, Utschig, Tiede and colleagues reported an alternative H2 -producing in vitro system using a self-assembled complex
of cyanobacterial PSI isolated from Synechococcus and cobaloxime
as the efficient proton reduction electrocatalysts (Utschig et al.,
2011a) in the presence of cyt c6 as a natural electron donor
to P700+ (see Fig. 6A). Cobaloximes are pseudomacrocyclic
bis(dimethylglyoxamato)cobalt complexes that were originally
developed as vitamin B12 alternative and discovered to perform
electrochemical proton reduction (Bakac et al., 1986; Razavet et al.,
2005). In this way, inexpensive and earth-abundant metal was used
for the first time as the proton reduction module operating in tandem with natural PSI as the photocatalytic module. Notably, the
hydrogen evolution rate for the PSI-cobaloxime hybrid complex
was shown to match that of the best reported for the cyt c6 -PSIPt nanoparticle hybrid system (246 ␮mol H2 mg Chl−1 h−1 ), when
recorded by the same authors under equivalent illumination conditions (Utschig et al., 2011a,b).
Another group of attractive molecular catalysts that has been
widely used in the biomimetic H2 -producing nanodevices are the
hydrogenase (H2 ase) enzymes, which can be linked with synthetic photosensitizers or PSI as the natural photocatalytic module.
H2 ases are generally divided into three independently evolved
classes termed [Fe]-, [FeFe]- and [NiFe]-H2 ases (Lubitz et al., 2007;
Vignais and Billoud, 2007; Fontecilla-Camps et al., 2007, 2009).
[FeFe]-H2 ases are found in bacteria and Eucarya, whereas [NiFe]H2 ases are found in bacteria and Archaea. A third type of H2 ases,
mononuclear [Fe]-H2 ase does not contain any iron–sulphur clusters and has been found only in some methanogenic Archaea.
H2 ases are the only molecular catalysts that are capable of
catalyzing both proton reduction and hydrogen oxidation with
efficiencies comparable to the platinum catalyst (Jones et al.,
2002). Only [NiFe]- and [FeFe]-H2 ases have been employed in
the biomimetic H2 -evolving systems, as the [Fe]- (also termed
Hmd) H2 ases are light sensitive, and thus, not useful in solar-tofuel biomimetic devices. [NiFe]- and [FeFe]-H2 ases are structurally
different, but catalyze the same reaction employing structurally
different catalytic sites (Fontecilla-Camps et al., 2009). Prokaryotic cyanobacteria usually employ [NiFe]-H2 ases, which are usually
less sensitive to irreversible inactivation by O2 . [FeFe]-H2 ases are
expressed in microalgae (e.g., C. reinhardtii) and anaerobic bacteria
(e.g., Chlamydomonas acetobutylicum) that produce H2 during fermentation or anaerobic respiration. The [FeFe]-H2 ases are rapidly
and often irreversibly inhibited by O2 , a feature that has hampered
the efficient use of these enzymes in biomimetic photocatalytic
nanodevices. The high-resolution X-ray structures of both types of
H2 ases revealed important features of the catalytic sites (Volbeda
et al., 1995, 1996; Peters et al., 1998; Nicolet et al., 1999; FontecillaCamps et al., 2007, 2009). Their active site is a bi-nuclear metal
complex based either on exclusively iron, or on a combination of
nickel and iron ions in a sulphur-rich environment, with Fe ions also
coordinated by cyanide and carbon monoxide (Ogata et al., 2002;
Fontecilla-Camps et al., 2009). Both catalytic sites usually include
multiple [4Fe–4S] clusters that participate in electron transfer
1647
reactions between the active site and the electron acceptor or donor
molecules that interact with the surface of the protein (Lubner et al.,
2010a).
Although [NiFe]-H2 ases are 10–100 times less biologically active
in H2 production compared to [FeFe]-H2 ases, the major advantage
of employing them in the solar-to-fuel biometric systems is their
oxygen tolerance. Additionally, these O2 -tolerant H2 ases are very
attractive candidates as the molecular catalysts of proton reduction
in the solid-state biomimetic photodevices, as discussed below.
In 2002, Rögner and colleagues proposed a model for a complete
photo-hydrogen producing nanodevice that would include PSI, PSII
and a H2 ase in a modular configuration, allowing for the combination of respective highly active proteins from various extremophilic
organisms. Such a modular system would be easily exchangeable
and should be separately characterized and optimized for all the
components (Wenk et al., 2002). As the first step toward the complete device, the authors reported optimization of the [NiFe]-H2 ase
from Thiocapsa roseopersicina. The isolated H2 ase was deposited
as Langmuir–Blodgett films on quartz glass or indium–tin oxide
(ITO) electrodes, and its H2 -evolving activity was measured with
respect to counter ions, presence of oxygen and the number of protein layers immobilized on an electrode (Wenk et al., 2002). The
authors demonstrated that poly-l-lysine and poly-butyl-viologen
as counter ions on the sub-phase stabilized the H2 ase on quartz
glass. The presence of Ca2+ , oxygen, excess amount or multiple
layers of protein on the surface of the electrode resulted in a significant loss of H2 production. The maximal hydrogen production
rate was achieved at 0.35 nmol H2 min−1 per monolayer of H2 ase
(Wenk et al., 2002).
The first example of a direct light-to-hydrogen conversion system using both H2 ase and PSI was reported in 2006 by the groups
of Okura and Friedrich (Ihara et al., 2006). The authors designed
an artificial fusion protein composed of the membrane-bound
[NiFe]-H2 ase (MBH) from the soil bacterium Ralstonia eutropha
and the PsaE extrinsic subunit of T. elongatus PSI. Both proteins
were connected by a small amino acid linker that replaced the
membrane-anchor at the C-terminus of the smallest H2 ase subunit,
HoxK. After spontaneous association with the PsaE-null mutant of
PSI, the resulting H2 ase-PsaE-PSI hybrid complex displayed lightdriven hydrogen production at a rate of 0.58 ␮mol H2 mg Chl−1 h−1 ,
in presence of ascorbic acid, dithiothreitol and tetramethyl phenylene diamine (TMPD) as the exogenous electron donors and
acceptors (Ihara et al., 2006). In this fusion complex, the most
distal [4Fe–4S] FB cluster of PSI was located approximately at a
14 Å distance to the distal accessory [4Fe–4S]-cluster of the H2 ase,
allowing for an efficient direct electron transfer between both ETCs.
A similar approach was used by Lenz, Rögner and colleagues, who
constructed a functional highly homogeneous hybrid complex of
the MBH-PsaE fusion protein and the PsaE-null mutant of Synechocystis PSI (Schwarze et al., 2010).
Ihara et al. (2006) discovered that such a hybrid complex
had the ability to interact with the native Fd, which acted as
a competitive electron acceptor, and therefore, also an inhibitor
of electron transport between PSI and the MBH H2 ase. As the
authors aimed at improving in vivo H2 production rates using a
hybrid complex of PSI and the MBH, they attempted to diminish the inhibitory effects of the Fd-dependent electron transfer
pathway. To this end, they engineered a fusion protein of PsaE
and cyt c3 from Desulfovibrio vulgaris (Ihara et al., 2006), which
(Amunts et al., 2010). Presented are the helices of several of the protein subunits, PsaA (yellow), PsaB (light turquoise), PsaC (dark blue), PsaD (light purple), PsaE (tangerine)
PsaF (black), PsaG (indigo), and the cofactors (color coding as in A, B, D, F). Other subunits and cofactors have been omitted for clarity. The views of PSI are approximately
with the threefold and twofold symmetry axis of the cyanoPSI and plant LHCI-PSI, respectively, in the membrane plane. The PDB coordinates are 1JB0 (cyanoPSI; Jordan
et al., 2001), 3LW5 (higher plant LHCI-PSI; Amunts et al., 2010), 2FRV and 1UBO ([NiFe]-hydrogenases; Volbeda et al., 1996; Ogata et al., 2002), 1FEH ([Fe]-hydrogenase;
Peters et al., 1998), 1C6S (cyt c6 ; Beissinger et al., 1998) and 9PCY (PC; Moore et al., 1991). Asc, ascorbate; DCPIP, 2,6-dichlorophenolindophenol; H2 ase, hydrogenase; MBH,
membrane-bound hydrogenase; MV, methyl viologen; MA, mercaptoacetic acid; PC, plastocyanin; PMS, N-methylphenazonium methyl sulphate.
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J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
is the natural electron donor for H2 ases in Desulfovibrio species
(Matias et al., 2001). This fusion protein was employed to reconstitute the activity of PSI lacking the PsaE subunit, resulting in a
hybrid complex that was able to interact with the specific H2 ase
while simultaneously exhibiting a decreased affinity for Fd (Ihara
et al., 2006).
A promising new approach toward efficient H2 photoproduction is based on the use of [FeFe]-H2 ases with a
natural affinity to Fd. Under anaerobiosis, several species of green
algae such as C. reinhardtii undergo hydrogen photo-production
catalyzed by a group of [FeFe]-H2 ases termed HydA. Happe and
colleagues demonstrated that Fd is critical for efficient electron
transport between PSI and HydA1 in Chlamydomonas by employing an elegant in vitro reconstitution system for H2 production
composed of PC (as a natural electron donor to P700+ ), LHCI-PSI
supercomplex, Fd (PetF protein) and a wild-type HydA1 H2 ase
(Winkler et al., 2009). Kinetic analyses of several site-directed
mutants of HydA1 and PetF allowed mapping of the key amino
acids essential for electrostatic interactions and electron transfer
between both proteins. In particular, a conserved HydA1 lysine
residue Lys396 seems to play a critical role for interaction with the
C-terminus of PetF, while the PetF residue Glu122 is essential for
docking the N-terminus of HydA1. The maximum H2 production
rate was reported at ∼90 ␮mol H2 mg Chl−1 h−1 (Winkler et al.,
2009). Although this yield is still in the range of in-solution systems, it demonstrates a potential of this O2 -sensitive [FeFe]-H2 ase
for the use in solar-to-hydrogen production systems.
One advantage of employing the diffusion-based H2 -producing
systems is the ability to vary and fine-tune the individual components without employing their specific chemical modifications
for immobilization within solid devices. A rather interesting variation of the diffusion-based biomimetic system for H2 production
has been reported, whereby hydrogen evolution was catalyzed by
the O2 -tolerant [NiFe]-H2 ase from the phototrophic purple sulphur bacterium T. roseopersicina operating in tandem with PSI from
cyanobacterium Synechocystis sp. PCC 6803. In this in vitro system,
H2 evolution was sustained in the presence of mercaptoacetic acid
(MA) as the sacrificial electron donor and methyl viologen (MV)
as the exogenous electron acceptor (and donor to the H2 ase), as
shown in Fig. 6B (Qian et al., 2008). This system evolved up to
0.5 ␮mol H2 mg Chl−1 h−1 . Despite such a low H2 evolution rate, the
system confirmed feasibility of a direct in-solution electron transfer between an exogenous electron donor, PSI and a catalytic centre
of the H2 ase.
A major advancement of the PSI-based H2 -producing in vitro and
solid-state systems has been achieved through application of the
“molecular wiring” technology developed by Golbeck, Bryant and
colleagues (Grimme et al., 2008, 2009; Lubner et al., 2010a,b, 2011).
This approach employs a “molecular wire” compound to connect
a terminal [4Fe–4S] cluster of PSI (FB cluster) directly to a H2 producing catalyst, which can be either the distal [4Fe–4S] cluster
of an [FeFe]- or [NiFe]-H2 ase (see Fig. 6D) or a noble metal nanoparticle (as depicted in Fig. 6E). The methodology involves constructing
mutant variants of both PSI and a H2 ase so that a molecular wire
can be attached to their surface-located iron–sulphur clusters. The
most distal surface-located iron–sulphur clusters in PSI and H2 ase
are each ligated by 4 cysteine residues, one of which can be altered
via site-directed mutagenesis to a glycine residue. These changes
expose iron atoms both in PSI and in H2 ase, allowing the –SH rescue
ligands from the molecular wire (usually a thiolate derivative) to
form two strong covalent disulphide bonds with the photocatalytic
(PSI) and proton-reducing (H2 ase or Pt/Au nanoparticle) modules.
Upon absorption of two photons by P700, the photoactivated electrons are efficiently transferred through the molecular wire from
PSI to the proton-reducing catalyst, which in turn reduces two protons to molecular hydrogen.
Very recently, molecular wiring of cyanobacterial PSI with the
[FeFe]-H2 ase from C. acetobutylicum allowed for light-induced H2
production at a spectacular rate of 2200 ␮mol mg Chl−1 h−1 , i.e. at
greater than two-fold electron throughput by this hybrid nanoconstruct compared to in vivo oxygenic photosynthesis (Lubner et al.,
2011). An important improvement of this technology was covalent
cross-linking of the natural electron donor, cyt c6 to the donor side
of PSI to ameliorate diffusion-based limitations of electron transfer
on the donor side (see Fig. 6D).
In contrast to high rates of H2 production when using
wired H2 ase as the proton reducing catalysts, molecular
wiring of the noble metal nanoparticles to the terminal FB
cluster of PSI resulted in rather modest rates of H2 production of 3.4 ␮mol H2 mg Chl−1 h−1 with a gold catalyst and
9.6 ␮mol H2 mg Chl−1 h−1 when platinum nanoparticles were used
(Grimme et al., 2008). Importantly, when cyt c6 was used as a
natural electron donor to the photooxidized dithol-wired PSI, the
H2 evolution yield increased 5-fold to 49.3 ␮mol H2 mg Chl−1 h−1
(Grimme et al., 2008), reiterating the importance of limiting the
diffusion-based donor side electron transfer to ensure higher rates
of hydrogen production.
PSI-based solid-state systems for H2 production
Hybrid biological/organic photochemical systems for H2
production
The approach of selective covalent modification of PSI
and its natural or synthetic electron donors and acceptors
overcomes the limitations of diffusion-based electron transfer processes. Evans et al. (2004) reported covalent linking of PC to
platinized PSI by employing a cross-linking reagent 1-ethyl-2-(3dimethylaminopropyl) carbodiimide hydrochloride, as depicted in
Fig. 6C. Compared to the diffusion-based setup, covalent linking of
PC and the donor side of PSI resulted in a 3-fold increased rate of
H2 evolution at an initial rate of 0.09 ␮mol H2 mg Chl−1 −1 (Evans
et al., 2004), demonstrating the importance of limiting diffusionbased electron transfer on the donor side for maximization of the
H2 production. The relatively low H2 production rate is most likely
due to the fact that diffusion limitation still applies to the reduction
of PC by Asc (sacrificial electron donor). Another possibility is that
at least a fraction of the spinach LHCI-PSI supercomplex used in
the study is likely to undergo photoinhibition or a fraction of Pt-PSI
conjugates may precipitate under experimental conditions applied
in the study, possibly leading to a significant loss of H2 production.
The hybrid systems described above have paved the way for the
development of immobilized PSI-bioconjugates within the solidstate systems for H2 production. The field is new, and so far a rather
limited number of efforts have employed such an attractive stateof-the art technology. To date, most in solido efforts have focused
on the application of photosynthetic complexes as photodetectors
or photovoltaic cells. For instance, Das et al. (2004) reported the
integration of spinach PSI and purple bacterial RC from Rhodobacter sphaeroides in solid state devices, creating photodetectors and
photovoltaic cells with internal quantum efficiencies of approximately 12%, an efficiency highly desirable for the purposes of H2
photo-evolution. They achieved a proper electronic integration of
the device by self-assembling an oriented monolayer of each photosynthetic RC. These were in turn stabilized with surfactant peptides
and coated with a protective organic semiconductor to ensure efficient electron flow between each half-cell.
Recently, several groups reported application of cyanobacterial
PSI as a photocathode that was molecular-wired to the protonreduction centre within the half-cell cathodic solid devices. Miyachi
et al. (2009) reported a novel molecular wire derived from vitamin
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
K1 (VK1 ) with a napthoquinone moiety that can connect to the FB
cluster of the T. elongatus PSI and a terpyridine moiety for connection to the Co (II) proton-reducing catalyst. A self-assembled
monolayer of PSI and the VK1 molecular wire was immobilized
on an indium tin oxide (ITO) electrode. This bioconjugate complex
showed the photocurrent action spectrum with a profile consistent to that of PSI, reflecting its suitability for H2 -production,
although H2 production itself was not investigated in this study
(Miyachi et al., 2009). Similarly, Terasaki et al. (2009) describe a
novel molecular ‘connector’ system, whereby an artificial wire (a
napthoquinone-viologen derivative NQC15 EV) assembled on a gold
electrode is plugged into one of the PSI redox active cofactors, the
A1 phylloquinone. The authors successfully obtained the desired
output of electrons from photo-activated PSI and demonstrated the
effectiveness of the molecular wiring technology for efficient coupling of PSI with the electrode by photoelectric transfer kinetics
analysis of the wire molecule (Terasaki et al., 2009).
Recently, Heberle and colleagues generated a H2 -evolving photoelectrode by immobilizing the PSI-[NiFe]-H2 ase hybrid complex
on the surface of a Ni-NTA-functionalized gold electrode, and orienting the donor side of PSI (through the His-tagged PsaF subunit)
at a close distance to the electrode (Krassen et al., 2009), as shown in
Fig. 6F. In this study, the full functionality of the PsaE-null mutant
of PSI from Synechocystis sp. PCC 6803 was rescued by binding a
fusion of the PsaE subunit and the membrane-bound/oxygen tolerant [NiFe]-H2 ase (MBH) from R. eutropha, a similar fusion complex
to that reported by Ihara et al. (2006). The estimated distance
between the FB cluster of PSI and the distal Fe-S cluster of the
MBH was 14–25 Å, indicating tight electronic coupling between
both ETCs in the engineered PSI-H2 ase hybrid complex. As a result,
more than 5000-fold enhanced light dependent H2 production
rate of 3000 ␮mol H2 mg Chl−1 h−1 was achieved. Unfortunately,
the assembly appears to be unstable, as half-life of the photocurrent was only 30 min. In addition, in order to maintain the reduced
state of the exogenous electron donor N-methylphenazonium
methyl sulphate (PMS), overpotential of −90 mV had to be applied.
Nonetheless, the system reflects the quintessential advantage of
solid state tightly coupled assemblies over diffusion-based systems.
Moreover, Krassen and colleagues clearly demonstrated that electron supply on the PSI donor side appears to be the major bottleneck
for this type of H2 -producing solid-state assemblies.
Future outlook
The two X-ray structures of cyanobacterial PSI and higher plant
LHCI-PSI supercomplex have revolutionized biophysical and biochemical studies on this molecular complex aimed at dissecting
the precise sequence of molecular events from light capture by the
antenna pigments through excitation of the P700 dimer, charge
separation and charge migration to the acceptor side of this complex. In particular, kinetic studies of electron transfer within PSI
and the discovery of two active branches of electron transfer in PSI
RCs have been aided tremendously by the detailed knowledge of
the exact ligand environment of the ETCs of PSI obtained through Xray crystallography combined with site-directed mutagenesis. Both
crystal structures have proven invaluable for probing interaction
with other components of the photosynthetic machinery during
adaptation responses to varying light conditions and stress factors. Nevertheless, a number of challenges remain to be addressed,
including dissecting the molecular regulatory pathways of CET
around PSI and its functional importance in higher plants, the exact
modes of interaction between PSI and the mobile LHCII antenna
during short-term adaptation of state transitions, and identification of the putative CET megasupercomplex in higher plants. The
biggest challenge of all is to obtain X-ray structures of extremely
1649
labile supercomplexes of PSI together with the other components
of photosynthetic machinery, such as the mobile LHCII antenna, Fd,
FNR and PC/cyt c6 .
There is no doubt that the inherent light-harvesting and
electron-transfer properties of natural PSI make this macromolecular complex particularly attractive for application in the
solar-to-fuel biomimetic devices reviewed in this article. The longlived charge-separated state P700+ FB − and an exceptionally low
redox potential associated with the distal FB cluster generate sufficient reducing power for production of molecular hydrogen in
man-made devices at neutral pH. While the H2 production yields
of the early semi-artificial systems could not compete with contemporary commercial hydrogen generation techniques, tremendous
progress in revealing the structure and function of PSI and H2 ases,
and a growing demand for a sustainable H2 source have led to a
recent revival of this research field. Nevertheless, this field is still
at the stage of experimental laboratory work and lacks any in-depth
knowledge regarding costs and large scale applications. Bruce and
colleagues argued that their self-assembling in vitro Pt-PSI-cyt c6
solar collector system could produce hydrogen with an energy yield
equivalent to that of 300 liters of gasoline per hectare per day. This
predicted yield would be more than one order of magnitude higher
than the gross yield of gasoline equivalents produced by contemporary agricultural biomass systems (Iwuchukwu et al., 2010). The
authors postulate that this system is capable of converting approximately 6% of solar radiation into usable fuel, the efficiency much
higher than that of the natural photosynthesis.
At present, it is difficult to predict with certainty the success of commercial-scale application of biomimetic solar-to-fuel
PSI-based devices. Nevertheless, some general aspects that affect
system efficiencies should be considered while engineering such
large-scale biomimetic devices. While systems based on noble
metal additives and those based on H2 ases exhibit comparable efficiencies, the major disadvantage is the high cost of noble metal
catalysts, as well as oxygen sensitivity of the most active H2 ases.
Understanding the precise kinetics of electron transfer within photosensitizer (PSI) module and a hydrogen-evolving catalyst (H2 ase
or synthetic proton reduction catalyst) and their inter-molecular
interactions is a prerequisite for the design of biomimetic hydrogen producing systems which have the potential to be economically
promising. Very recently, the significant interest in utilizing the
highly stable natural PSI for generation of hydrogen as a clean
solar fuel has resulted in engineering vastly improved biomimetic
devices with some of the highest light-driven turnover yields of
H2 ever observed. The pioneering work by Golbeck, Bryant and
colleagues on development of the molecular wiring technology,
reviewed in this article, emphasizes the importance of minimizing
energy losses due to diffusion-based electron transfer within solarto-fuel devices. Similarly, tight electronic coupling between the
terminal FB cluster of PSI and the distal Fe–S cluster of the oxygentolerant H2 ase in a solid-state system, such as the system developed
by Heberle, Lenz and colleagues (Krassen et al., 2009), represents
an attractive approach for improved solar-to-fuel devices. Thus, the
critical issue of amelioration of losses due to the donor and acceptor side rate limitation around PSI has been successfully addressed
in small-scale solid-state and in vitro systems.
We firmly believe that the next few years will bring considerable technological advances that will lead to construction of
the complete highly stable and electronically coupled solid-state
H2 -evolving biomimetic device utilizing PSI as a photocathodic
module which, in conjunction with the biological or synthetic
proton reducing centre, will use electrons and protons produced
through photooxidation of water in the anodic half-cell for production of molecular hydrogen at a high yield. In this way, electron
diffusion obstacles and artificial donor/acceptor availability will
be overcome. Undoubtedly, the biggest challenge of all will be to
1650
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
ensure redox integration of both half-cells into a highly efficient
solar-to-hydrogen device, utilizing much broader light spectrum
than the photosynthetically active radiation (PAR) used by the natural antenna systems and photochemical RCs of PSI and PSII. To this
end, efforts are being made to construct solid-state heterojunction
tandem devices that will use spectroscopically separated artificial antenna systems both within the anode and cathode half-cells
capturing light wavelengths below and above photosynthetically
active radiation.
Acknowledgement
We gratefully acknowledge funding from the Polish Ministry
of Science and Higher Education (MNiSW grant no. 844/N-ESFEuroSolarFuels/10/2011/0).
References
Allen AE, Laroche J, Maheswari U, Lommer M, Schauer N, Lopez PJ, et al. Whole-cell
response of the pennate diatom Phaeodactylum tricornutum to iron starvation.
Proc Natl Acad Sci USA 2008;105:10438–43.
Allen JP, Fever G, Yeates TO, Komiya H, Rees DC. Structure of the reaction center from Rhodobacter sphaeroides R-26: the cofactors. Proc Natl Acad Sci USA
1987;84:5730–4.
Alric J. Cyclic electron flow around photosystem I in unicellular green algae. Photosynth Res 2010;106:47–56.
Amunts A, Nelson N. Plant photosystem I design in the light of evolution. Structure
2009;17:637–50.
Amunts A, Drory O, Nelson N. The structure of plant photosystem I supercomplex at
3.4 Å. Nature 2007;447:58–63.
Amunts A, Toporik H, Borovikova A, Nelson N. Structure determination and
improved model of plant photosystem I. J Biol Chem 2010;285:3478–85.
Andrizhiyevskaya EG, Schwabe TM, Germano M, D’Haene S, Kruip J, Van Grondelle
R. Spectroscopic properties of PSI-IsiA supercomplexes from the cyanobacterium
Synechococcus PCC 7942. Biochim Biophys Acta 2002;556:265–72.
Antonkine ML, Jordan P, Fromme P, Krauss N, Golbeck JH, Stehlik D. Assembly of
protein subunits within the stromal ridge of photosystem I. Structural changes
between unbound and sequentially PS I-bound polypeptides and correlated
changes of the magnetic properties of the terminal iron–sulfur clusters. J Mol
Biol 2003;327:671–97.
Archer MD, Barber J. Photosynthesis and photoconversion. In: Archer MD, Barber
J, editors. Molecular to global photosynthesis. London: Imperial College Press;
2004. p. 1–41.
Armbrust EV, Berges JA, Bowler C, Green BR, Martinez D, Putnam NH, et al.
The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and
metabolism. Science 2004;306:79–86.
Bakac A, Brynildson ME, Espenson JH. Characterization of the structure, properties, and reactivity of a cobalt (II) macrocyclic complex. Inorg Chem
1986;25:4108–14.
Baniulis D, Yamashita E, Whitelegge JP, Zatsman AI, Hendrich MP, Hasan SS,
et al. Structure–function, stability, and chemical modification of the cyanobacterial cytochrome b6 f complex from Nostoc sp. PCC 7120. J Biol Chem
2009;284:9861–9.
Barber J, Nield J, Morris EP, Hankamer B. Subunit positioning in photosystem II
revisited. Trends Biochem Sci 1999;24:43–5.
Barber J, Nield J, Duncan J, Bibby TS. Accessory chlorophyll proteins in cyanobacterial photosystem I. In: Golbeck GH, editor. Photosystem I: the light-driven
plastocyanin:ferredoxin oxidoreductase. Dordrecht: Springer; 2006. p. 99–117.
Bautista JA, Rappaport F, Guergova-Kuras M, Cohen RO, Golbeck JH, Wang JY, et al.
Biochemical and biophysical characterization of photosystem I from phytoene
desaturase and zeta-carotene desaturase deletion mutants of Synechocystis sp.
PCC 6803: evidence for PsaA- and PsaB-side electron transport in cyanobacteria.
J Biol Chem 2005;280:20030–41.
Beissinger M, Sticht H, Sutter M, Ejchart A, Haehnel W, Rosch P. Solution structure of
cytochrome c6 from the thermophilic bacterium Synechococcus elongatus. EMBO
J 1998;17:26–36.
Ben-Shem A, Frolow F, Nelson N. Crystal structure of plant photosystem I. Nature
2003;426:630–5.
Ben-Shem A, Frolow F, Nelson N. Evolution of photosystem I-from symmetry
through pseudo-symmetry to asymmetry. FEBS Lett 2004;564:274–80.
Bibby TS, Nield J, Barber J. Iron deficiency induces the formation of an antenna ring
around trimeric photosystem I in cyanobacteria. Nature 2001a;412:743–5.
Bibby TS, Nield J, Barber J. Three-dimensional model and characterization of the iron
stress-induced CP43 -photosystem I supercomplex isolated from the cyanobacterium Synechocystis PCC 6803. J Biol Chem 2001b;276:43246–52.
Bibby TS, Nield J, Partensky F, Barber J. Oxyphotobacteria. Antenna ring around
photosystem I. Nature 2001c;413:590.
Bibby TS, Mary I, Nield J, Partensky F, Barber J. Low-light-adapted Prochlorococcus species possess specific antennae for each photosystem. Nature
2003a;424:1051–4.
Bibby TS, Nield J, Chen M, Larkum AWD, Barber J. Structure of a photosystem II supercomplex isolated from Prochloron didemni retaining its
chlorophyll a/b light-harvesting system. Proc Natl Acad Sci USA 2003b;100:
9050–4.
Blankenship RE. Molecular mechanisms of photosynthesis. Oxford: Blackwell Science; 2002.
Blankenship RE. Early evolution of photosynthesis. Plant Physiol 2010;154:434–8.
Blankenship RE, Tiede DM, Barber J, Brudvig GW, Fleming G, Ghirardi M, et al.
Comparing photosynthetic and photovoltaic efficiencies and recognizing the
potential for improvement. Science 2011;332:805–9.
Boekema EJ, Hifney A, Yakushevska AE, Piotrowski M, Keegstra W, Berry S, et al. A
giant chlorophyll–protein complex induced by iron deficiency in cyanobacteria.
Nature 2001;412:745–8.
Bottin H, Hanley J, Lagoutte B. Role of acidic amino acid residues of PsaD subunit
on limiting the affinity of photosystem I for ferredoxin. Biochem Biophys Res
Commun 2001;287:833–6.
Brudvig GW. Water oxidation chemistry of photosystem II. Phil Trans R Soc Lond B
2008;363:1211–9.
Busch A, Hippler M. The structure and function of eukaryotic photosystem I. Biochim
Biophys Acta 2011;1807:864–77.
Busch A, Nield J, Hippler M. The composition and structure of photosystem Iassociated antenna from Cyanidioschyzon merolae. Plant J 2010;62:886–97.
Chauhan D, Folea IM, Jolley CC, Kouril R, Lubner CE, Lin S, et al. A novel photosynthetic strategy for adaptation to low-iron aquatic environments. Biochemistry
2011;50:686–92.
Chen M, Blankenship RE. Expanding the solar spectrum used by photosynthesis.
Trends Plant Sci 2011;16:427–31.
Chitnis VP, Chitnis PR. PsaL subunit is required for the formation of photosystem I trimers in the cyanobacterium Synechocystis sp. PCC 6803. FEBS Lett
1993;336:330–4.
Cogdell RJ, Gardiner AT, Hashimoto H, Brotosudarmo TH. A comparative look at
the first few milliseconds of the light reactions of photosynthesis. Photochem
Photobiol Sci 2008;7:1150–8.
Collins A, Wen J, Blankenship R. Photosynthetic light-harvesting complexes. In:
Wydrzynski T, Hillier W, editors. Molecular solar fuels. London: RSC Publishing;
2011. p. 85–106.
Croce R, Dorra D, Holzwarth AR, Jennings RC. Fluorescence decay and spectral
evolution in intact photosystem I of higher plants. Biochemistry 2000;39:
6341–8.
Croce R, Morosinotto T, Castelletti S, Breton J, Bassi R. The Lhca antenna complexes
of higher plants photosystem I. Biochim Biophys Acta 2002;1556:29–40.
Das R, Kiley PJ, Segal M, Norville J, Yu AA, Wang L, et al. Integration of photosynthetic protein molecular complexes in solid-state electronic devices. Nano Lett
2004;4:1079–83.
Dashdorj N, Xu W, Cohen RO, Golbeck JH, Savikhin S. Asymmetric electron transfer in cyanobacterial Photosystem I: charge separation and secondary electron
transfer dynamics of mutations near the primary electron acceptor A0 . Biophys
J 2005;88:1238–49.
Deisenhofer J, Epp O, Sinning I, Michel H. Crystallographic refinement at 2.3 Å
resolution and refined model of the photosynthetic reaction centre from
Rhodopseudomonas viridis. J Mol Biol 1995;246:429–57.
DeLano WL. The PyMOL molecular graphics system. San Carlos, CA, USA: DeLano
Scientific; 2002, http://www.pymol.org.
Deng Z, Aliverti A, Zanetti G, Arakaki AK, Ottado J, Orellano EG, et al. A productive NADP+ binding mode of ferredoxin-NADP+ reductase revealed by protein
engineering and crystallographic studies. Nat Struct Biol 1999;6:847–53.
Desquilbet TE, Duval JC, Robert B, Houmard J, Thomas JC. In the unicellular red alga
Rhodella violacea iron deficiency induces an accumulation of uncoupled LHC.
Plant Cell Physiol 2003;44:1141–51.
Diner BA, Rappaport F. Structure, dynamics, and energetics of the primary photochemistry of photosystem II of oxygenic photosynthesis. Annu Rev Plant Biol
2002;53:551–71.
Doan JM, Schoefs B, Ruban AV, Etienne AL. Changes in the LHCI aggregation state
during iron repletion in the unicellular red alga Rhodella violacea. FEBS Lett
2003;533:59–62.
Drop B, Webber-Birungi M, Fusetti F, Kouril R, Redding KE, Boekema EJ, et al.
of Chlamydomonas reinhardtii contains nine light-harvesting Ccmplexes (Lhca)
located on one side of the core. J Biol Chem 2011;286:44878–87.
Durnford DG, Deane JA, Tan S, McFadden GI, Gantt E, Green BR. A phylogenetic assessment of the eukaryotic light-harvesting antenna proteins, with implications for
plastid evolution. J Mol Evol 1999;48:59–68.
El-Mohsnawy E, Kopczak MJ, Schlodder E, Nowaczyk M, Meyer HE, Warscheid B, et al.
Structure and function of intact photosystem I monomers from the cyanobacterium Thermosynechococcus elongatus. Biochemistry 2010;49:4740–51.
Evans BR, O’Neill HM, Hutchens SA, Bruce BD, Greenbaum E. Enhanced photocatalytic hydrogen evolution by covalent attachment of plastocyanin to
photosystem I. Nano Lett 2004;4:1815–9.
Evans MCV, Cammack R. The effect of the redox state of the bound iron–sulphur
centres in spinach chloroplasts on the reversibility of P700 photooxidation at
low temperatures. Biochem Biophys Res Commun 1975;63:187–93.
Evans MCV, Sihra CK, Cammack R. The properties of the primary electron acceptor
in the Photosystem I reaction centre of spinach chloroplasts and its interaction with P700 and the bound ferredoxin in various oxidation–reduction states.
Biochem J 1976;158:71–7.
Ferreira KN, Iverson TM, Maghlaoui K, Barber J, Iwata S. Architecture of the photosynthetic oxygen-evolving center. Science 2004;303:1831–8.
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
Fischer N, Sétif P, Rochaix JD. Site-directed mutagenesis of the PsaC subunit of photosystem I. FB is the cluster interacting with soluble ferredoxin. J Biol Chem
1999;274:23333–40.
Fischer N, Hippler M, Sétif P, Jacquot JP, Rochaix JD. The PsaC subunit of photosystem
I provides an essential lysine residue for fast electron transfer to ferredoxin.
EMBO J 1998;17:849.
Fontecilla-Camps JC, Volbeda A, Cavazza C, Nicolet Y. Structure/function relationships of [NiFe]- and [FeFe]-hydrogenases. Chem Rev 2007;107:4273–430.
Fontecilla-Camps JC, Amara P, Cavazza C, Nicolet Y, Volbeda A. Structure–function
relationships of anaerobic gas-processing metalloenzymes. Nature
2009;460:814–22.
Frazao C, Soares CM, Carrondo MA, Pohl E, Dauter Z, Wilson KS, et al. Ab initio
determination of the crystal structure of cytochrome c6 and comparison with
plastocyanin. Structure 1995;3:1159–69.
Ganeteg U, Klimmek F, Jansson S. Lhca5—an LHC-type protein associated with photosystem I. Plant Mol Biol 2004;54:641–51.
Gardian Z, Bumba L, Schrofel A, Herbstova M, Nebesarova J, Vacha F. Organisation of photosystem I and photosystem II in red alga Cyanidium caldarium:
encounter of cyanobacterial and higher plant concepts. Biochim Biophys Acta
2007;1767:725–31.
Germano M, Yakushevska AE, Keegstra W, Van Gorkom HJ, Dekker JP, Boekema EJ.
Supramolecular organization of photosystem I and light-harvesting complex I
in Chlamydomonas reinhardtii. FEBS Lett 2002;525:121–5.
Glazer AN. Light harvesting by phycobilisomes. Annu Rev Biophys Biophys Chem
1985;14:47–77.
Golbeck JH, McDermott AE, Jones WK, Kurtz DM. Evidence for the existence of
[2Fe–2S] as well as [4Fe–4S] clusters among FA , FB and Fx. Implications for
the structure of the Photosystem I reaction center. Biochim Biophys Acta
1987;891:94–8.
Grabolle M, Dau H. Energetics of primary and secondary electron transfer
in photosystem II membrane particles of spinach revisited on basis of
recombination-fluorescence measurements. Biochim Biophys Acta 2005;1708:
209–18.
Green BR, Durnfold DG. The chlorophyll-carotenoid proteins of oxygenic photosynthesis. Annu Rev Plant Physiol Plant Mol Biol 1996;47:685–714.
Greenbaum E. Platinized chloroplasts: a novel photocatalytic material. Science
1985;230:1373–5.
Grimme RA, Lubner CE, Bryant DA, Golbeck JH. Photosystem I/molecular wire/metal
nanoparticle bioconjugates for the photocatalytic production of H2 . J Am Chem
Soc 2008;130:6308–9.
Grimme RA, Lubner CE, Golbeck JH. Maximizing H2 production in Photosystem
I/dithiol molecular wire/platinum nanoparticle bioconjugates. Dalton Trans
2009;45:10106–13.
Grotjohann I, Fromme P. Structure of cyanobacterial photosystem I. Photosynth Res
2005;85:51–72.
Guergova-Kuras M, Boudreaux B, Joliot A, Joliot P, Redding K. Evidence for two
active branches for electron transfer in photosystem I. Proc Natl Acad Sci USA
2001;98:4437–42.
Guskov A, Kern J, Gabdulkhakov A, Broser M, Zouni A, Saenger W. Cyanobacterial
photosystem II at 2.9-Å resolution and the role of quinones, lipids, channels and
chloride. Nat Struct Mol Biol 2009;16:334–42.
Guss JM, Freeman HC. Structure of oxidized poplar plastocyanin at 1.6 Å resolution.
J Mol Biol 1983;169:521–63.
Haldrup A, Naver H, Scheller HV. The interaction between plastocyanin and photosystem I is inefficient in transgenic Arabidopsis plants lacking the PSI-N subunit
of photosystem I. Plant J 1999;17:689–98.
Haldrup A, Simpson DJ, Scheller HV. Down-regulation of the PSI-F subunit of
photosystem I (PSI) in Arabidopsis thaliana. The PSI-F subunit is essential for
photoautotrophic growth and contributes to antenna function. J Biol Chem
2000;275:31211–8.
Haumann M, Liebisch P, Müller C, Barra M, Grabolle M, Dau H. Photosynthetic O2 formation tracked by time-resolved x-ray experiments. Science
2005;310:1019–21.
Hippler M, Drepper F, Haehnel W, Rochaix JD. The N-terminal domain of PsaF: precise recognition site for binding and fast electron transfer from cytochrome c6
and plastocyanin to photosystem I of Chlamydomonas reinhardtii. Proc Natl Acad
Sci USA 1998;95:7339–44.
Hippler M, Drepper F, Rochaix JD, Muhlenhoff U. Insertion of the N-terminal part
of PsaF from Chlamydomonas reinhardtii into photosystem I from Synechococcus
elongatus enables efficient binding of algal plastocyanin and cytochrome c6 . J
Biol Chem 1999;274:4180–8.
Hippler M, Reichert J, Sutter M, Zak E, Altschmied L, Schröer U, et al. The plastocyanin
binding domain of photosystem I. EMBO J 1996;15:6374–84.
Hohmann-Marriott MF, Blankenship RE. Evolution of photosynthesis. Annu Rev
Plant Biol 2011;62:515–48.
Holzwarth AR, Müller MG, Niklas J, Lubitz W. Ultrafast transient absorption studies
on photosystem I reaction centers from Chlamydomonas reinhardtii. 2. Mutations
near the P700 reaction center chlorophylls provide new insight into the nature
of the primary electron donor. Biophys J 2006;90:552–65.
Hu Q, Miyashita H, Iwasaki I, Kurano N, Miyachi S, Iwaki M, et al. A photosystem I
reaction center driven by chlorophyll d in oxygenic photosynthesis. Proc Natl
Acad Sci USA 1998;95:13319–23.
Ihalainen JA, Jensen PE, Haldrup A, Van Stokkum IH, Van Grondelle R, Scheller HV,
et al. Pigment organization and energy transfer dynamics in isolated photosystem I (PSI) complexes from Arabidopsis thaliana depleted of the PSI-G, PSI-K,
PSI-L, or PSI-N subunit. Biophys J 2002;83:2190–1.
1651
Ihara M, Nishihara H, Yoon KS, Lenz O, Friedrich B, Nakamoto K, et al. Light-driven
hydrogen production by a hybrid complex of a [NiFe]-hydrogenase and the
cyanobacterial photosystem I. Photochem Photobiol 2006;82:676–82.
Iwuchukwu IJ, Vaughn M, Myers N, O’Neill H, Frymier P, Bruce BD. Self-organized
photosynthetic nanoparticle for cell-free hydrogen production. Nat Nanotechnol 2010;5:73–9.
Jennings RC, Zucchelli G, Croce R, Garlaschi FM. The photochemical trapping rate
from red spectral states in PSI-LHCI is determined by thermal activation of
energy transfer to bulk chlorophylls. Biochim Biophys Acta 2003;1557:91–8.
Jensen PE, Bassi R, Boekema EJ, Dekker JP, Jansson S, Leister D, et al. Structure, function and regulation of plant photosystem I. Biochim Biophys Acta
2007;1767:335–52.
Johnson GN. Physiology of PSI cyclic electron transport in higher plants. Biochim
Biophys Acta 2011;1807:384–9.
Joliot P, Joliot A. In vivo analysis of the electron transfer within photosystem I: are
the two phylloquinones involved? Biochemistry 1999;38:11130–6.
Jones AK, Sillery E, Albracht SPJ, Armstrong FA. Direct comparison of the electrocatalytic oxidation of hydrogen by an enzyme and a platinum catalyst. Chem
Commun 2002;8:866–7.
Jordan P, Fromme P, Witt HT, Klukas O, Saenger W, Krauss N. Threedimensional structure of cyanobacterial photosystem I at 2.5 Å resolution.
Nature 2001;411:909–17.
Juhas M, Büchel C. Properties of photosystem I antenna protein complexes of the
diatom Cyclotella meneghiniana. J Exp Bot; http://dx.doi.org/10.1093/jxb/ers049,
in press.
Kameda H, Hirabayashi K, Wada K, Fukuyama K. Mapping of protein–protein interaction sites in the plant-type [2Fe–2S] ferredoxin. PLoS One 2011;6:e21947.
Kamiya N, Shen JR. Crystal structure of oxygen-evolving photosystem II from
Thermosynechococcus vulcanus at 3.7-Å resolution. Proc Natl Acad Sci USA
2003;100:98–103.
Karapetyan NV, Dorra D, Schweitzer G, Bezsmertnaya IN, Holzwarth AR. Fluorescence spectroscopy of the longwave chlorophylls in trimeric and monomeric
photosystem I core complexes from the cyanobacterium Spirulina platensis. Biochemistry 1999;36:13830–7.
Kargul J, Barber J. Photosynthetic acclimation: structural reorganisation of light harvesting antenna—role of redox-dependent phosphorylation of major and minor
chlorophyll a/b binding proteins. FEBS J 2008;275:1056–68.
Kargul J, Barber J. Structure and function of photosynthetic reaction centres. In:
Wydrzynski T, Hillier W, editors. Molecular solar fuels. London: RSC Publishing;
2011. p. 107–42.
Kargul J, Nield J, Barber J. Three-dimensional reconstruction of a light-harvesting
complex I-photosystem I (LHCI-PSI) supercomplex from the green alga Chlamydomonas reinhardtii, Insights into light harvesting for PSI. J Biol Chem
2003;278:16135–41.
Kargul J, Turkina MV, Nield J, Benson S, Vener AV, Barber J. Light-harvesting complex
II protein CP29 binds to photosystem I of Chlamydomonas reinhardtii under State
2 conditions. FEBS J 2005;272:4797–806.
Karplus PA, Daniels MJ, Herriott JR. Atomic structure of ferredoxin-NADP+ reductase:
prototype for a structurally novel flavoenzyme family. Science 1991;251:60–6.
Kerfeld CA, Anwar HP, Interrante R, Merchant S, Yeates TO. The structure of chloroplast cytochrome c6 at 1.9 Å resolution: evidence for functional oligomerization.
J Mol Biol 1995;250:627–47.
Knoetzel J, Mant A, Haldrup A, Jensen PE, Scheller HV. PSI-O, a new 10-kDa subunit
of eukaryotic photosystem I. FEBS Lett 2002;510:145–8.
Kouril R, Arteni AA, Lax J, Yeremenko N, D’Haene S, Rögner M, et al. Structure and
functional role of supercomplexes of IsiA and photosystem I in cyanobacterial
photosynthesis. FEBS Lett 2005a;579:3253–7.
Kouril R, Zygadlo A, Arteni AA, de Wit CD, Dekker JP, Jensen PE, et al. Structural
characterization of a complex of photosystem I and light-harvesting complex II
of Arabidopsis thaliana. Biochemistry 2005b;44:10935–40.
Kramer DM, Avenson TJ, Edwards GE. Dynamic flexibility in the light reactions of
photosynthesis governed by both electron and proton transfer reactions. Trends
Plant Sci 2004;9:349–57.
Krassen H, Schwarze A, Friedrich B, Ataka K, Lenz O, Heberle J. Photosynthetic hydrogen production by a hybrid complex of photosystem I and [NiFe]-Hydrogenase.
ACS Nano 2009;3:4055–61.
Kruip J, Bald D, Boekema EJ, Rögner M. Evidence for the existence of trimeric
and monomeric photosystem I complexes in the thylakoid membranes from
cyanobacteria. Photosynth Res 1994;40:279–86.
Kurisu G, Zhang H, Smith JL, Cramer WA. Structure of the cytochrome b6 f complex
of oxygenic photosynthesis: tuning the cavity. Science 2003;302:1009–14.
Kurisu G, Kusunoki M, Katoh E, Yamazaki T, Teshima K, Onda Y, et al. Structure
of the electron transfer complex between ferredoxin and ferredoxin-NADP(+)
reductase. Nat Struct Biol 2001;8:117–21.
La Roche J, Van der Staay GWM, Partensky F, Ducret A, Aebersold R, Li R, et al.
Independent evolution of the prochlorophyte and green plant chlorophyll a/b
light-harvesting proteins. Proc Natl Acad Sci USA 1996;93:15244–8.
Lee JW, Collins RT, Greenbaum E. Molecular ionic probes: a new class of Hill reagents
and their potential for nanofabrication and biometallocatalysis. Phys Chem B
1998;102:2095–100.
Lee JW, Tevault CV, Blankinship SL, Collins RT, Greenbaum E. Photosynthetic water
splitting: in situ photoprecipitation of metallocatalysts for photoevolution of
hydrogen and oxygen. Energy Fuels 1994;8:770–3.
Li Y, Van der Est A, Lucas MG, Ramesh VM, Gu F, Petrenko A, et al. Directing electron
transfer within photosystem I by breaking H-bonds in the cofactor branches.
Proc Natl Acad Sci USA 2006;103:2144–9.
1652
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
Liu Z, Yan H, Wang K, Kuang T, Zhang J, Gui L, et al. Crystal structure of spinach major
light-harvesting complex at 2.72 Å resolution. Nature 2004;428:287–92.
Loll B, Kern J, Saenger W, Zouni A, Biesiadka J. Towards complete cofactor
arrangement in the 3.0 Å resolution structure of photosystem II. Nature
2005;438:1040–4.
Lubitz W, Reijerse E, van Gastel M. [NiFe] and [FeFe] hydrogenases studied by advanced magnetic resonance techniques. ACS Chem Rev 2007;107:
4331–65.
Lubner CE, Grimme R, Bryant DA, Golbeck JH. Wiring photosystem I for direct solar
hydrogen production. Biochemistry 2010a;49:404–14.
Lubner CE, Knörzer P, Silva PJN, Vincent KA, Happe T, Bryant DA, et al. Wiring an
[FeFe]-hydrogenase with photosystem I for light-induced hydrogen production.
Biochemistry 2010b;49:10264–6.
Lubner CE, Applegate AM, Knorzer P, Ganago A, Bryant DA, Happe T, et al. Solar
hydrogen-producing bionanodevice outperforms natural photosynthesis. Proc
Natl Acad Sci USA 2011;108:20988–91.
Lucinski R, Schmid VH, Jansson S, Klimmek. Lhca5 interaction with plant photosystem I. FEBS Lett 2006;580:6485–8.
Lunde C, Jensen PE, Haldrup A, Knoetzel J, Scheller HV. The PSI-H subunit of photosystem I is essential for state transitions in plant photosynthesis. Nature
2000;408:613–5.
MacColl R. Allophycocyanin and energy transfer. Biochim Biophys Acta
2004;1657:73–81.
Matias PM, Soares CM, Saraiva LM, Coelho R, Morais J, Gall JL, et al. [NiFe]
hydrogenase from Desulfovibrio desulfuricans ATCC 27774: gene sequencing,
three-dimensional structure determination and refinement at 1.8 Å and modelling studies of its interaction with the tetrahaem cytochrome c3 . J Biol Inorg
Chem 2001;6:63–81.
Matsuzaki M, Misumi O, Shin IT, Maruyama S, Takahara M, Miyagishima SY, et al.
Genome sequence of the ultrasmall unicellular red alga Cyanidioschyzon merolae
10D. Nature 2004;428:653–7.
McDermott AE, Yachandra VK, Guiles RD, Sauer K, Klein MP, Parrett KG. EXAFS structural study of FX , the low-potential Fe-S center in photosystem I. Biochemistry
1989;28:8056–9.
Melkozernov AN, Bibby TS, Lin S, Barber J, Blankenship RE. Time-resolved absorption
and emission show that the CP43 antenna ring of iron-stressed Synechocystis
sp. PCC6803 is efficiently coupled to the photosystem I reaction centre core.
Biochemistry 2003;42:3893–903.
Melkozernov AN, Kargul J, Lin S, Barber J, Blankenship RE. Energy coupling in the
PSI-LHCI supercomplex from the green alga Chlamydomonas reinhardtii. J Phys
Chem B 2004;108:10547–55.
Melkozernov AN, Kargul J, Lin S, Barber J, Blankenship RE. Spectral and kinetic analysis of the energy coupling in the PS I-LHC I supercomplex from the green alga
Chlamydomonas reinhardtii at 77 K. Photosynth Res 2005;86:203–15.
Millsaps JF, Bruce BD, Lee JW, Greenbaum E. Nanoscale photosynthesis: photocatalytic production of hydrogen by platinized photosystem I reaction centers.
Photochem Photobiol 2001;73:630–5.
Miyachi M, Yamanoi Y, Yonezawa T, Nishihara H, Iwai M, Inoue Y. Surface immobilization of PSI using vitamin K1-like molecular wires for fabrication of a
bio-photoelectrode. J Nanosci Nanotechnol 2009;9:1722–6.
Miyashita H, Ikemoto H, Kurano N, Adachi K, Chihara M, Miyachi S. Chlorophyll d as
a major pigment. Nature 1996;383:402.
Moore JM, Lepre CA, Gippert GP, Chazin WJ, Case DA, Wright PE. High-resolution
structure of reduced French bean plastocyanin and comparison with the crystal
structure of poplar plastocyanin. J Mol Biol 1991;221:533–55.
Morales R, Charon MH, Hudry-Clergeon G, Petillot Y, Norager S, Medina M, et al.
Refined X-ray structures of the oxidized, at 1.3 Å, and reduced, at 1.17 Å, [2Fe–2S]
ferredoxin from the cyanobacterium Anabaena PCC7119 show redox-linked conformational changes. Biochemistry 1999;38:15764–73.
Moreland JL, Gramada A, Buzko OV, Zhang Q, Bourne PE. The Molecular Biology
Toolkit (MBT): a modular platform for developing molecular visualization applications. BMC Bioinformatics 2005;6:21.
Moseley JL, Allinger T, Herzog S, Hoerth P, et al. Adaptation to Fe-deficiency requires
remodeling of the photosynthetic apparatus. EMBO J 2002;21:6709–20.
Müller MG, Slavov C, Luthra R, Redding KE, Holzwarth AR. Independent initiation
of primary electron transfer in the two branches of the photosystem I reaction
center. Proc Natl Acad Sci USA 2010;107:4123–8.
Muraki N, Seo D, Shiba T, Sakurai T, Kurisu G. Asymmetric dimeric structure of
ferredoxin-NAD(P)+ oxidoreductase from the green sulfur bacterium Chlorobaculum tepidum: implications for binding ferredoxin and NADP+ . J Mol Biol
2010;401:403–14.
Murray JW, Duncan J, Barber J. CP43-like chlorophyll binding proteins: structural
and evolutionary implications. Trends Plant Sci 2006;11:152–8.
Naumann B, Stauber EJ, Busch A, Sommer F, Hippler M. N-terminal processing
of Lhca3 Is a key step in remodeling of the photosystem I-light-harvesting
complex under iron deficiency in Chlamydomonas reinhardtii. J Biol Chem
2005;280:20431–41.
Naumann B, Busch A, Allmer J, Ostendorf E, Zeller M, Kirchhoff H, et al. Comparative quantitative proteomics to investigate the remodeling of bioenergetic
pathways under iron deficiency in Chlamydomonas reinhardtii. Proteomics
2007;7:3964–79.
Neilson JA, Durnford DG. Structural and functional diversification of the
light-harvesting complexes in photosynthetic eukaryotes. Photosynth Res
2010;106:57–71.
Nelson N. Plant photosystem I-the most efficient nano-photochemical machine. J
Nanosci Nanotechnol 2009;9:1709–13.
Nelson N. Photosystems and global effects of oxygenic photosynthesis. Biochim
Biophys Acta 2011;1807:856–63.
Nelson N, Ben-Shem A. The structure of photosystem I and evolution of photosynthesis. BioEssays 2005;27:914–22.
Nicolet Y, Piras C, Legrand P, Hatchikian CE, Fontecilla-Camps JC. Desulfovibrio
desulfuricans iron hydrogenase: the structure shows unusual coordination to
an active site Fe binuclear center. Struct Fold Des 1999;7:13–23.
Nield J, Morris EP, Bibby TS, Barber J. Structural analysis of the photosystem
I supercomplex of cyanobacteria induced by iron deficiency. Biochemistry
2003;42:3180–8.
Ogata H, Mizoguchi Y, Mizuno N, Adachi S, Yasouka N, Yagi T, et al. Structural studies
of the carbon monoxide complex of [NiFe]-hydrogenase from Desulfobivri vulgaris: suggestion for the initial activation site for dihydrogen. J Am Chem Soc
2002;124:11628–35.
Pan X, Li M, Wan T, Wang L, Jia C, Hou Z, et al. Structural insights into energy
regulation of light-harvesting complex CP29 from spinach. Nat Struct Mol Biol
2011;18:309–15.
Peltier G, Tolleter D, Billon E, Cournac L. Auxiliary electron transport pathways in
chloroplasts of microalgae. Photosynth Res 2010;106:19–31.
Peng L, Shikanai T. Supercomplex formation with photosystem I is required for the
stabilization of the chloroplast NADH dehydrogenase-like complex in Arabidopsis. Plant Physiol 2011;155:1629–39.
Peng L, Shimizu H, Shikanai T. The chloroplast NAD(P)H dehydrogenase complex
interacts with photosystem I in Arabidopsis. J Biol Chem 2008;283:34873–9.
Peng L, Fukao Y, Fujiwara M, Takami T, Shikanai T. Efficient operation of NAD(P)H
dehydrogenase requires supercomplex formation with photosystem I via minor
LHCI in Arabidopsis. Plant Cell 2009;21:3623–40.
Peters JW, Lanzilotta WN, Lemon BJ, Seefeldt LC. X-ray crystal structure of the Fe-only
hydrogenase (Cpl) from Clostridium pasteurianum to 1.8 Å resolution. Science
1998;282:1853–8.
Poluektov OG, Paschenko SV, Utschig LM, Lakshmi KV, Thurnauer MC. Bidirectional
electron transfer in photosystem I: direct evidence from high-frequency timeresolved EPR spectroscopy. J Am Chem Soc 2005;127:11910–1.
Qian DJ, Liu AR, Nakamura C, Wenk SO, Miyake J. Photoinduced hydrogen evolution
in an artificial system containing photosystem I, hydrogenase, methyl viologen
and mercaptoacetic acid. Chin Chem Lett 2008;19:607–10.
Ramesh VM, Gibasiewicz K, Lin S, Bingham SE, Webber AN. Bidirectional electron
transfer in photosystem I: accumulation of A0 - in A-side or B-sidemutants of the
axial ligand to chlorophyll A0 . Biochemistry 2004;43:1369–75.
Rappaport F, Guergova-Kuras M, Nixon PJ, Diner BA, Lavergne J. Kinetics
and pathways of charge recombination in photosystem II. Biochemistry
2002;41:8518–27.
Razavet M, Artero V, Fontecave M. Proton electroreduction catalyzed
by cobaloximes: functional models for hydrogenases. Inorg Chem
2005;44:4786–95.
Redinbo MR, Cascio D, Choukair MK, Rice D, Merchant S, Yeates TO. The 1.5-Å crystal structure of plastocyanin from the green alga Chlamydomonas reinhardtii.
Biochemistry 1993;32:10560–7.
Rochaix J-D. Regulation of photosynthetic electron transport. Biochim Biophys Acta
2011;1807:375–83.
Rögner M, Muhlenhoff U, Boekema EJ, Witt HT. Mono-, di- and trimeric PSI reaction
center complexes isolated from the thermophilic cyanobacterium Synechococcus
sp. size, shape and activity. Biochim Biophys Acta 1990;1015:415–24.
Ruban AV, Johnson MP. Xanthophylls as modulators of membrane protein function.
Arch Biochem Biophys 2010;504:78–85.
Ruffle SV, Mustafa AO, Kitmitto A, Holzenburg A, Ford RC. The location of the
mobile electron carrier ferredoxin in vascular plant photosystem I. J Biol Chem
2000;275:36250–5.
Sadekar S, Raymond J, Blankenship RE. Conservation of distantly related membrane
proteins: photosynthetic reaction centers share a common structural core. Mol
Biol Evol 2006;23:2001–7.
Santabarbara S, Galuppini L, Casazza AP. Bidirectional electron transfer in the reaction centre of photosystem I. J Integr Plant Biol 2010;52:735–49.
Santabarbara S, Jasaitis A, Byrdin M, Gu F, Rappaport F, Redding K. Additive effect of
mutations affecting the rate of phylloquinone reoxidation and directionality of
electron transfer within photosystem I. Photochem Photobiol 2008;84:1381–7.
Santabarbara S, Kuprov I, Fairclough WV, Purton S, Hore PJ, Heathcote P, et al.
Bidirectional electron transfer in photosystem I: determination of two distances between P700+ and A1 − in spin-correlated radical pairs. Biochemistry
2005;44:2119–28.
Scheller HV, Jensen PE, Haldrup A, Lunde C, Knoetzel J. Role of subunits in eukaryotic
photosystem I. Biochim Biophys Acta 2001;1507:41–60.
Schubert WD, Klukas O, Saenger W, Witt HT, Fromme P, Krauss NJ. A common ancestor for oxygenic and anoxygenic photosynthetic systems: a comparison based
on the structural model of photosystem I. J Mol Biol 1998;280:297–314.
Schwarze A, Kopczak MJ, Rögner M, Lenz O. Requirements for construction of a functional hybrid complex of photosystem I and [NiFe]-hydrogenase. Appl Environ
Microbiol 2010;76:2641–51.
Serre L, Vellieux FM, Medina M, Gomez-Moreno C, Fontecilla-Camps JC, Frey M.
X-ray structure of the ferredoxin:NADP+ reductase from the cyanobacterium
Anabaena PCC 7119 at 1.8 Å resolution, and crystallographic studies of NADP+
binding at 2.25 Å resolution. J Mol Biol 1996;263:20–39.
Sétif P, Fischer N, Lagoutte B, Bottin H, Rochaix JD. The ferredoxin docking site of
photosystem I. Biochim Biophys Acta 2002;1555:204–9.
Shikanai T. Cyclic electron transport around photosystem I: genetic approaches.
Annu Rev Plant Biol 2007;58:199–217.
J. Kargul et al. / Journal of Plant Physiology 169 (2012) 1639–1653
Siegbahn PEM. Recent theoretical studies of water oxidation for photosystem II. J
Photochem Photobiol B 2011;104:94–9.
Slavov C, Ballottari M, Morosinotto T, Bassi R, Holzwarth AR. Trap-limited charge
separation kinetics in higher plant photosystem I complexes. Biophys J
2008;94:3601–12.
Sproviero EM, Gascón JA, McEvoy JP, Brudvig GW, Batista VS. Computational studies
of the O2 -evolving complex of photosystem II and biomimetic oxomanganese
complexes. Coord Chem Rev 2008;252:395–415.
Standfuss J, Terwisscha van Scheltinga AC, Lamborghini M, Kuhlbrandt W.
Mechanisms of photoprotection and nonphotochemical quenching in pea lightharvesting complex at 2.5 Å resolution. EMBO J 2005;24:919–28.
Stauber EJ, Busch A, Naumann B, Svatos A, Hippler M. Proteotypic profiling of LHCI
from Chlamydomonas reinhardtii provides new insights into structure and function of the complex. Proteomics 2009:398–408.
Storf S, Stauber EJ, Hippler M, Schmid VH. Proteomic analysis of the photosystem
I light-harvesting antenna in tomato (Lycopersicon esculentum). Biochemistry
2004;43:9214–24.
Storf S, Jansson S, Schmid VH. Pigment binding, fluorescence properties, and
oligomerization behavior of Lhca5, a novel light-harvesting protein. J Biol Chem
2005;280:5163–8.
Stroebel D, Choquet Y, Popot JL, Picot D. An atypical haem of the cytochrome b6 f
complex. Nature 2003;426:413–8.
Tejero J, Martinez-Julvez M, Mayoral T, Luquita A, Sanz-Aparicio J, Hermoso
JA, et al. Involvement of the pyrophosphate and the 2 -phosphate binding
regions of ferredoxin-NADP+ reductase in coenzyme specificity. J Biol Chem
2003;278:49203–14.
Terasaki N, Yamamoto N, Hiraga T, Yamanoi Y, Yonezawa T, Nishihara H, et al.
Plugging a molecular wire into photosystem I: reconstitution of the photoelectric conversion system on a gold electrode. Angew Chem Int Ed Engl
2009;48:1585–7.
Thangaraj B, Jolley CC, Sarrou I, Bultema JB, Greyslak J, Whitelegge J, et al. Efficient
light harvesting in a dark, hot, acidic environment: the structure and function
of PSI-LHCI from Galdieria sulphuraria. Biophys J 2011;100:135–43.
Tomo T, Okubo T, Akimoto S, Yokono M, Miyashita H, Tsuchiya T, et al. Identification
of the special pair of photosystem II in a chlorophyll d-dominated cyanobacterium. Proc Natl Acad Sci USA 2007;104:7283–8.
Umena YK, Kawakami K, Shen JR, Kamiya N. Crystal structure of oxygen-evolving
photosystem II at a resolution of 1.9 Å. Nature 2011;473:55–60.
Utschig LM, Silver CS, Mulfort KL, Tiede DM. Nature-driven photochemistry for
catalytic solar hydrogen production: a photosystem I-transition metal catalyst
hybrid. J Am Chem Soc 2011a;133:16334–7.
Utschig LM, Dimitrijevic NM, Poluektov OG, Chemerisov SD, Mulfort KL, Tiede DM.
Photocatalytic hydrogen production from noncovalent biohybrid photosystem
I/Pt nanoparticle complexes. J Phys Chem Lett 2011b;2:236–41.
1653
Vanselow C, Weber AP, Krause K, Fromme P. Genetic analysis of the photosystem I subunits from the red alga, Galdieria sulphuraria. Biochim Biophys Acta
2009;1787:46–59.
Veith T, Büchel C. The monomeric photosystem I-complex of the diatom
Phaeodactylum tricornutum binds specific fucoxanthin chlorophyll proteins
(FCPs) as light-harvesting complexes. Biochim Biophys Acta 2007;1767:
1428–35.
Veith T, Brauns J, Weisheit W, Mittag M, Büchel C. Identification of a specific
fucoxanthin-chlorophyll protein in the light harvesting complex of photosystem I in the diatom Cyclotella meneghiniana. Biochim Biophys Acta
2009;1787:905–12.
Vignais PM, Billoud B. Occurrence, classification, and biological function of hydrogenases: an overview. Chem Rev 2007;107:4206–72.
Volbeda AM, Charon H, Piras C, Hatchikian EC, Frey M, Fontecilla-Camps JC.
Crystal structure of the [NiFe] hydrogenase from Desulfovibrio gigas. Nature
1995;373:580–7.
Volbeda AM, Garcin E, Piras C, DeLacey AL, Fernandez VM, Hatchikan EC, et al. Structure of the [NiFe] hydrogenase active site: evidence for biologically uncommon
Fe ligands. J Am Chem Soc 1996;118:12989–96.
Watanabe M, Kubota H, Wada H, Narikawa R, Ikeuchi M. Novel supercomplex organization of photosystem I in Anabaena and Cyanophora paradoxa. Plant Cell Physiol
2011;52:162–8.
Watanabe T, Kobayashi M, Hongu A, Nakazato M, Hiyama T. Evidence, that a
chlorophyll a dimer constitutes the photochemical reaction centre I (P700) in
photosynthetic apparatus. FEBS Lett 1985;235:252–6.
Wenk SO, Qian DJ, Wakayama T, Nakamura C, Zorin N, Rögner M, et al. Biomolecular device for photoinduced hydrogen production. Int J Hydrogen Energy
2002;27:1489–93.
Winkler M, Kuhlgert S, Hippler M, Happe T. Characterization of the key step
for light-driven hydrogen evolution in green algae. J Biol Chem 2009;284:
36620–7.
Yamashita E, Zhang H, Cramer WA. Structure of the cytochrome b6 f complex:
quinone analogue inhibitors as ligands of heme cn. J Mol Biol 2007;370:
39–52.
Yan J, Kurisu G, Cramer WA. Structure of the cytochrome b6 f complex: binding
site and intraprotein transfer of the quinone analogue inhibitor 2,5-dibromo3-methyl-6-isopropyl-p-benzoquinone. Proc Natl Acad Sci USA 2006;103:
67–74.
Yeremenko N, Kouril R, Ihalainen JA, D’Haene S, Van Oosterwijk N, Andrizhiyevskaya
EG, et al. Supramolecular organization and dual function of the IsiA chlorophyllbinding protein in cyanobacteria. Biochemistry 2004;43:10308–13.
Zouni A, Witt HT, Kern J, Fromme P, Krauss N, Saenger W, et al. Crystal structure of photosystem II from Synechococcus elongatus at 3.8 Å resolution. Nature
2001;409:739–43.