Doctoral Thesis / Dissertation, 2010
I.I. Evolutionary Origin of Oxygenic Photosynthesis
I.II. Basic Features of Chloroplasts
I.III. Architecture of Oxygenic Photosynthesis
I.III.I. Molecular Architecture
I.III.II. Alternative PSI Dependant Electron Pathways
I.III.III. Reaction Centers 9 IIIIIV Antenna Systems
I.III.IV.I. Light-Harvesting Pigments
I.IV. Structural Biology of Photosystems
I.IV.I. Crystallization and Structure Determination
I.IV.I.I. Crystallization of Membrane Proteins 23 IV General Structural Resemblance between PSI and PSII 26 IVI Photosystem II
I.VI.I. Macroscopic Architecture of the Core Complex
I.VI.I.I. Arrangement of Core Subunits
I.VI.I.II. Interactions Between the Monomers 30 IVIII Additional Membrane-extrinsic Subunits 30 IVIIII Functional Site - Catalytic Role of PSII
I.VI.III.I. Electron Transport Chain and Charge Separation
I.VI.III.II. Water Oxidation Complex (WOC)
I.VI.IV. Plant PSII Antenna - Light Harvesting Complex II (LHCII)
I.VI.V. Remodeling PSII Antenna as Acclimation Response to Light-driven Stimuli 36 IVII Photosystem I
I.VII.I. Macroscopic Architecture of the Core Complex
I.VII.I.I. Arrangement of Subunits
I.VII.I.II. Interactions between the Monomers in Cyanobacteria versus Plant Monomers
I.VII.II. Binding of Soluble Electron Carriers
I.VII.III. Functional Site - Catalytic Role of PSI
I.VII.V. Plant PSI Antenna - Light Harvesting Complex I (LHCI)
I.VII.VI. Remodeling PSI Antenna as Acclimation Response to Environmental Stimuli
II. SUPPLEMENTARY METHODS
II.I. Crystal Cryoprotection
II.II. Data Collection
II.II.I. Quantitative factors - slicing method, partially recorded reflections and oscillation range considerations
II.II.II. Qualitative factors - exposure time, overloads and estimated uncertainties
II.III. Data Processing
II.IV. Molecular Replacement
II.V. Model Building and Refinement
III.I. Paper Review
III.II. Amunts A, Drory O and Nelson N (2007) The structure of a plant Photosystem I supercomplex at 34 Å resolution Nature 447, 58-63
III.III. Amunts A and Nelson N (2009) Plant Photosystem I design in the light of evolution Structure 17, 637-650
III.IV. Amunts A, Toporik H, Borovikova A and Nelson N (2010) Structure determination and improved model of plant Photosystem I Journal of Biological Chemistry 285, 3478-3486
III.V. Amunts A, Ben-Shem A and Nelson N (2005) Solving the structure of plant Photosystem I - biochemistry is vital Photochem Photobiol Science 4(12), 1011-1015
IV.I. Strategies Employed in the Interpretation of the X-ray Diffraction Data
IV.I.I. Molecular Replacement of Plant PSI
IV.I.II. Considerations for Low Resolution Structure Determination
IV.II. Functional Importance of Plant PSI Unique Subunits
IV.III. Ferredoxin Mediated Electron Transfer in PSI
IV.IV. Chlorophylls and Carotenoids: Beyond Orientation
IV.IV.I. Overall Composition
IV.IV.II. Unique Pigment Positions Affected by Selection Pressures
IV.IV.III. 102 + 61 > 163 ?
IV.IV.III.I. Are Gap Pigments Unique to PSI-LHCI ?
IV.V. Structural Organization of Plant PSI Antenna System - LHCI
V. TOWARD THE NEXT QUANTUM LEAP
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We are dwarves on giants ’ backs and for this reason we can look far. - Bernard of Chartres (12th century), philosopher.
Photosynthesis is a highly complex, multistep biological process by which solar energy is absorbed and converted into chemical energy needed to power life. It is almost the sole process by which the chemical energy to maintain living organisms is generated and therefore it is a primary engine of the biosphere. Thus, this remarkable process not only provides the foundation for life on the Earth, but also has profoundly altered our planet over the geological time scale [Falkowski et al., 1998; Blankenship, 2001; Lane, 2002; Xiong et al., 2002; Archer and Barber, 2004].
Oxygenic photosynthesis began soon after life itself, about 3.5 billion years ago, when first aquatic photosynthetic bacteria, cyanobacteria, have appeared in the depth of large ancient water reservoirs, and the onset of oxygen took place [Schopf, 1983; Woese, 1987; Schopf and Packer, 1987; Olson and Pierson, 1987; Schopf, 1993; Nisbet et al., 1995; Mulkidjanian and Junge, 1997; Nisbet and Fowler, 1999; Rasmussen, 2000; Schopf, 2000; Farquhar et al., 2000; Des Marais, 2001; Brasier et al., 2002; Clausen and Junge, 2004; Rashby et al., 2007; Barber, 2008; Larkum, 2008]. In the terms of energy production, the photosynthetic process had an advantage of a few orders of magnitude in size over the available oxidation-reduction reactions associated with weathering and hydrothermal activity [Olson et al., 1985; Martin et al., 1987; Elderfield and Schultz, 1996; De Marais, 1997]. Moreover, the ability of primordial oxygenic photosynthetic bacteria to reduce NADP+ to NADPH for organic biosynthesis by cleaving water made this evolutionary innovation extremely powerful [Buick, 1992; Brocks et al., 1999; Summons et al., 1999; Dismukes et al., 2001]. It freed life from its sole dependence upon abiotic chemical sources of reducing power, such as hydrothermal sources and weathering [De Marais, 2000]. Using water as a universal reductant, photosynthesis also had an infinite source of electrons. In addition, these cyanobacteria enjoyed anaerobic conditions that protected them from singlet oxygen damage, and relied upon wide water shielding from excess light intensities [Xiong and Bauer, 2002; Nelson and Ben Shem, 2005]. This, in conjunction with constant supplies of just enough sunlight, essential moisture, and abundant nutrients, resulted in massive production of organic carbon and the flourishing of photosynthetic communities in the distant geological past [De Marais, 2000].
However, it should be emphasized that Dismukes et al., (2001) proposed that bicarbonate preceded water as a thermodynamically preferred reductant for oxygenic photosynthesis in phototrophic bacteria with no living analogues yet discovered. The thermodynamic argument for an original bicarbonate substrate for oxygenic photosynthesis remains unchallenged. Importantly, the firstly emerged photosynthetic organisms also involved metabolism thermostability and temperature constraints [Schwartzman and Lineweaver, 2004; Schwartzman et al., 2008].
In any case, a cardinal side effect of the photosynthetic advance was the enormous increase in the production of oxygen, which was progressively released and accumulated in the atmosphere of the early Earth, turning the atmosphere into an oxidizing rather than a reducing ambient and transforming the geochemistry of the planet [Falkowski and Godfrey, 2008]. Oxygen buildup and consequent formation of insoluble iron-oxide (Fe2O3), which depleted oceans and lithosphere from iron, initiated extreme environmental changes that triggered what is known as the "Big Bang of Evolution" [Barber, 2004], a sequence of drastic evolutionary events that included sweeping diversification of O2-dependent life. Many of multiple effects and feedbacks of the appearance of free diatomic oxygen in Earth’s early atmosphere such as influence on the evolution of lipids, secondary metabolites and the selection of trace elements that are critical for maintaining electron traffic within organisms are not yet fully understood [Anbar and Knoll, 2002; Quigg et al., 2003; Raymond and Segre`, 2006]. However, it is clear that oxygen buildup led to the formation of a stratospheric ozone layer that absorbed UV radiation [Farquhar et al., 2000] and to the emergence of two of the most significant biological innovation about 2.2 - 1.8 billion years ago. First, the occurrence of an aerobic respiration, which was able to harness a more powerful metabolic energy source, because free molecular oxygen allowed a much greater thermodynamic efficiency in the oxidation of organic matter [Falkowski and Godfrey, 2008]. Second, the development of further advanced, but primarily energy-inefficient, Eucarya domain [Moreira and Lopez-Garcia, 1998; Hatman and Federov, 2002; Knoll, 2003; Javaux, 2007].
Later on, sometime in between the late Proterozoic era and the early Cambrian period (~ 1 billion years ago), a single-celled protest of the eukaryotic life forms (host) engulfed and retained a free-living primordial photosynthetic cyanobacterium through concomitant endosymbiosis [Margulis and Fester, 1991; Gray, 1992; Bhattacharya and Medlin, 1995; Douglas, 1998; Delwiche, 1999; Gray, 1999; Cavalier-Smith, 2000; McFadden, 2001; Palmer, 2003; Bhattacharya et al., 2004]. Over time, the metabolic systems of the two cells gradually became coordinated and the prokaryote was reduced to a double membrane-bound plastid and vertically transmitted to subsequent generations [Reyes-Prieto et al., 2007]. This resulted in the formation of chloroplasts, special Plantae ancestor organelles in which the process of oxygenic photosynthesis occurs [Gray, 1999; Besendahl et al., 2000; Yoon et al., 2004, Dyall et al., 2004; recently reviewed by Gould et al., 2008 and Keeling, 2010].
The bacterial-like staining properties of chloroplasts were originally recognized in the end of nineteenth century by microscopists J. Sachs (1882), A. Schimper (1883) and R. Altmann (1890). About 20 years later, C. Mereschkowsky (1905) culminated these observations into the hypothesize that chloroplasts are derived from cyanobacteria. L. Margulis later formalized the Theory of Endosymbiosis (1971), which posits that plastids and mitochondria of eukaryotic cells derive from bacterial endosymbionts.
The emergence of chloroplasts enabled communities of primitive algae to inhabit the surface of marine areas near the shore. Supplemented by nearer ground oxygen presence, it subsequently allowed the surface of the Earth to be habitable by plants approximately 500 million years ago [Grambast, 1974; Lewis and McCourt, 2004; recently reviewed by Sørensen, 2010]. Plants evolved quickly on land and were so successful that although the land surface of the Earth forms only ~30% of the total area, the land supplies over 50% of the photosynthetic products on the planet [Falkowski and Raven, 1997]. This success of plants supplied herbivorous animals with a diverse source of food, and gave the final crowning of the tree of life, opening up the continents to the wealth of diverse ecosystems known today.
Chloroplasts are lens-shaped, dynamic organelles, which move within the plant cell to maximize light absorption [Seng, 1908; Zurzycki, 1955] and to minimize photo- damage [Zurzycki, 1957]. Their dynamic nature is something at which to marvel. The flux of energy through the photosynthetic pathway goes from zero at predawn to a daily maximum at the sun's zenith and then back to zero at dusk. There is no other organelle that has such a huge range of metabolic activity over the course of a day. A plant cell contains normally up to 100 chloroplasts [recently reviewed by Rebeiz, 2010]. Each chloroplast is delineated by a double envelope membrane, which sometimes appears to circulate around the main body of the organelle [recently reviewed by Taiz and Zeiger, 2010]. Chloroplasts also contain an additional membrane system, the internal photosynthetic membrane, known as the thylakoids, first observed by W. Menke (1960, 1962) in electron micrographs of chloroplast cross sections. This membrane continuum of flattened vesicles stacks is characterized by a massive folding [Shimoni et al., 2005; Mustardy et al., 2008]. Essentially, at the chloroplast level modulation, it is the thylakoid membrane derived metabolic signals that operate the acclimatized responses [Anderson et al., 1995; Walters et al., 2003]. The thylakoid membrane is defined by 3 distinct domains: grana - the cylindrical compactly stacked structures; stroma lamellae - non-stacked double membrane regions, which connect adjacent grana; grana margins - include specific protein composition, different from the grana [Wollenberger et al, 1994; Kaftan et al., 2002]. The fluid compartment that surrounds the thylakoids and where the carbon metabolism reactions are sited, is known as the stroma, while the aqueous interior inside the thylakoids is defined as the lumen.
In 1937, R. Hill used ferricyanide as an artificial electron acceptor to discover that isolated from leaves chloroplasts are able to evolve oxygen. Today, we know that thylakoid membranes provide exclusive sites of light absorption and the primary reactions that transform light energy into biochemical energy, by hosting all the photosynthetic pigments, electron-transport complexes and the ATP synthase [Whatley et al., 1963]. The thylakoid membrane also catalyzes many biochemical processes and faces the destructive effect of reactive singlet oxygen. In addition, it has a relatively high protein:lipid ratio and the intrinsic complexes are closely packed, which makes it the most complicated membrane in nature [Vothknecht and Westhoff, 2001; recently reviewed by Wada and Murata, 2009]. Importantly, the four membrane complexes that drive the light reaction are not evenly distributed over the thylakoid. PSI and ATP synthase localizes mainly to the stroma lamella, segregated from PSII, which is almost exclusively found in the grana, whereas cytochrome b6f preferentially populates the grana and the grana margins [Anderson, 1999; Albertsson, 2001; Anderson, 2002; Kaftan et al., 2002; Kim et al., 2005; Daum et al., 2010].
Oxygenic photosynthesis is accomplished by a series of reactions driven by the four multisubunit membrane complexes (Figure 1): Photosystem II (PSII), cytochrome b 6 f, Photosystem I (PSI) and ATP synthase [Hill and Bendall, 1960; Duysens et al., 1961; Mitchell, 1961; McCarty and Racker, 1966; Jagendorf, 1967; Cramer and Butler; 1967; reviewed by Nelson and Ben Shem, 2004]. In the current section, some of the general properties of the four complexes are discussed, while a more profound analysis of PSII and PSI is presented in section I.VI. and I.VII. respectively.
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Figure 1 - The components of the light reactions in oxygenic photosynthesis.
(A) Structural models of protein components of photosynthetic membrane in the order they appear in the electron transport chain and described in the text. (B) The corresponding functional components of the electron transport chain. PSI and PSII are represented by one monomer each. The dimeric cytochrome b 6 f is reduced to one functional half. Arrows represent electrons pathways through the system. The magenta arrows represent direct electron transport, whereas the green arrows indicate the transport of an electron together with a proton, that is, in the form of a reduced plastoquinone.
PSII is defined as water-plastoquinone oxidoreductase. PSII oxidizes water (the electron donor for the whole process) into four protons (H+) and molecular oxygen (O2), catalyzing the light-driven electron transport to a quinone (QB) [recently reviewed by Guskov et al., 2010]. The four protons are released into the lumen and contribute to the establishment of the pH across the membrane. Two protons access the QB binding pocket from the stromal side of the thylakoid membrane and the reduced plastoquinol PQH2 leaves PSII through an exit channel into the membrane. It is subsequently replaced by a PQ from the quinone pool. The plastoquinol PQH2 is hydrophobic and therefore is able to diffuse through the membrane to the following complex, the cytochrome b 6 f, which functionally couples the light driven electron transfer events of PSI and PSII.
Cytochrome b 6 f uses the Q-cycle mechanism to couple the electron transfer with proton translocation and the generation of a transmembrane potential postulated by P. Mitchell (1961, 1976). Following the PQH2 binding to the cytochrome b 6 f hydrophobic pocket, close to the luminal side of the membrane, the two H+ are released into the lumen. One electron is funneled through the Q-cycle (similar to that guided by the respiration chain homologue, the bc 1 complex), which involves heme b L, heme b H, additional covalently bound heme c and plastoquinone bound on the stromal face of the membrane for taking up protons from the stroma [Berry et al., 2000; Cramer et al., 1996]. The other electron is further diverted to the 2Fe2S cluster and then to cytochrome f (heme f), which is located at the luminal membrane extrinsic domain of the b 6 f complex [Zhang et al., 1998; Kurisu et al., 2003; Stroebel et al., 2003]. The cytochrome f subunit contains a docking site for the soluble electron carriers plastocyanin and cytochrome c6, which transfer the electrons further to PSI [Cramer et al., 1996].
The reduced electron carriers dock at a luminal idention of PSI, in close proximity to the oxidized co-factors of the PSI electron transport chain (special pair, discussed below). The binding mode of reduced plastocyanin and cytochrome c6 varies between different organisms, from a weak diffusion controlled interaction in some cyanobacteria to the formation of a tight complex in higher plants [Haehnel et al., 1994; Hippler et al., 1997; De la Cerda et al., 1999; Chitnis, 2001; Durán et al., 2004].
PSI catalyzes the light-driven electron transfer from plastocyanin/cytochrome c6 (donor) at the luminal side of the membrane to ferredoxin/flavodoxin (acceptor) at the stromal side of the membrane [recently reviewed by Amunts and Nelson, 2009a]. The donor binding to PSI initiates the translocation of an electron across the membrane through a chain of cofactors consisting of two pairs of chlorophylls, a phylloquinone and three 4Fe4S clusters. From the terminal 4Fe4S cluster the electron is transferred to ferredoxin or flavodoxin, docked at the stromal hump of the complex. These proteins transfer the electron to ferredoxin NADP+ reductase (FNR) for the reduction of NADP+ to NADPH [Arnon 1951; Vishniac and Ochoa, 1951; Tolmach, 1951; Shin et al., 1963; Hall and Rao, 1977; Hauska and Trebst, 1977]. Overall, the electron transfer reactions lead to the formation of an electrochemical proton gradient across the thylakoid membrane, which is used for the synthesis of ATP from ADP and Pi by the ATP synthase. [Arnon et al., 1954; Arnon, 1955; Trebst, 1974; Thornber, 1975; Goldbeck et al., 1977; Junge, 1977; Williams, 1977; Trebst, 1978; Avron, 1981; Malkin, 1981; Codgell, 1983].
In the dark reactions, photosynthetically generated high energy products ATP and reduced NADPH are used in a variety of metabolic processes, which take place mainly but not exclusively in chloroplasts, including CO2 assimilation in the BensonCalvin cycle [Bassham et al., 1954; Bassham and Calvin, 1957], nitrate metabolism, lipid, amino acids, glucose and pigment synthesi and the modulation of gene expression [Ort and Yocum, 1996; Nelson and Yocum, 2006].
Electron flow from PSI is a complex and dynamic process, which has been intensively studied, but yet poorly understood [Eberhard et al., 2008; Iwai et al., 2010; Alric, 2010]. In addition to the major photosynthetic electron transfer (usually related as linear electron transfer) described above, different abiotic and biotic stress conditions may lead to electron redirection from PSI toward alternative electron sinks. Possible diversions include the b 6 f complex, the quinone pool, monodehydroascorbate radical (MDA), shunting electrons into the respiratory chain and more.
The stimuli/motivation which induce these branched pathways are of three main folds: (i) Balancing the ATP/NADPH stoichiometry. Diverting electrons from the reducing side of PSI back to the Q pool or b 6 f complex (usually related as cyclic electron transfer, since electrons generated on the reducing side of PSI can be reinjected at its donor side), first discovered by D. Arnon (1959), creates an alternative electron flow that contributes to the generation of an extra pH without net production of reducing equivalents [Johnson, 2005; Muneckage et al., 2002]. This is important because the assimilation of CO2 in the Benson-Calvin cycle requires ATP:NADPH ratio of 3:2 [Allen, 2002], which is not satisfied by linear electron transfer [Kramer et al., 2004]. However, there has been a long debate as to what extent the cyclic electron transfer exists under physiological conditions [Joliot and Joliot, 2006; Rumeau et al., 2007; Shikanai, 2007] and the actual pathway by which electrons are transferred back from the donor side of PSI is still under debate [Bendall and Manasse, 1995; Albertsson, 2001; Peltier and Cournac, 2002; Joliot and Joliot, 2002; Stroebel et al., 2003; Golding et al., 2004; Nandha et al., 2007; Eberhard et al., 2008; Dal Corso et al., 2008].
(ii) Photoprotection. Absorption of light in excess of the capacity of the linear electron transfer may cause photodamage [Heber and Walker, 1992; Horton et al., 1996]. The existence of alternative sinks actively counterbalances the full reduction of the photosynthetic electron carriers under the low light conditions when the Benson- Calvin cycle is deactivated. This way the branched pathways allow direct adjustment of electron utilization and therefore provide a further element of flexibility to the capacity of the electron transfer process. In addition, the photodamage may be induced by O2 oxidation of the reduced ferredoxin, at the PSI or PSII acceptor side, known as Mehler reaction [Badger et al., 2000]. The reaction leads to the production of O·2, which is rapidly converted to H2O2 by a superoxide dismutase (SOD) [Asada, 1999; Rizhsky et al., 2003]. A photoprotective mechanism suggests that O·2 is consumed by ascorbate peroxidase, which synthesizes a monodehydroascorbate radical (MDA) from ascorbate and H2O2 [Miyake and Asada, 1992]. MDA can consequently be efficiently reduced directly by photosynthetic electrons at a rate comparable to that of NADP+ reduction [Forti and Elli, 1995].
(iii) Fueling other metabolic pathways/networks. NADPH provides the reducing power for many metabolic pathways [Edwards and Walker, 1983; Geigenberger et al., 2005] and triggers regulatory networks controlled by the thioredoxin/peroxyredoxin systems [Buchanan and Balmer, 2005]. In addition, NADPH generated by the photosynthetic electron transfer can be reoxidized by the mitochondrial respiratory chain through the exchange of reducing power between the two organelles [Scheibe, 1987; Rebeille and Gans, 1988; Bulte et al., 1990; Cardol et al., 2003; Noctor et al., 2004; Eberhard et al., 2008]. Therefore, the acceptor side of PSI is a key site for integration of photosynthesis into the cell metabolism.
All types of photosynthetic systems are constructed from two principle components: reaction center (RC) and light-harvesting antenna (see next section). In the RCs the photochemical reaction takes place, the conversion of photoelectric energy to electrochemical potential. This process involves the movement of electrical charge across the membrane, illustrated in Figure 1. There are two types of RC, known as Type I and Type II, and briefly described below. Despite having a possible common evolutionary origin [Deisenhofer et al., 1985; Schubert et al., 1998 ; Rhee et al., 1998; Baymann et al., 2001; Heathcote et al., 2002], the two types differ in the key thermodynamic properties, linked mainly to the identity of the terminal electron acceptor.
Within the heart of both types RCs, two closely spaced pigment molecules, (bacterio)chlorophylls, are located, known as the special pair and given the symbol "P" for pigment. Due to the unique location and the physico-chemical environment provided by the protein moiety, the special pair forms an efficient trap for excitation energy. Thus, P is promoted to an excited electronic state, the primary radical P*, which provides an extremely strong reducing specie [Arnon, 1989; Krall and Edwards, 1992; Chitnis, 1996]. The generation of P* occurs either by direct photon absorption or, more commonly, by energy transfer from the antenna system at very high quantum efficiencies.
Once P* is formed, the stage is set for the primary reaction of photosynthesis, which conducts a charge separation. P* operates as a reducing agent and primary electron donor for the following series of acceptors (electron transport chain) that transfer the electron outside the RC. At the first step, P* rapidly loses an electron to a nearby electron acceptor molecule (A), generating an ionpair state P+A-, thus photoelectric excitation energy has been transformed to chemical redox driving force. The primary electron acceptor "A" can be either (bacterio)chlorophyll (Type I RC) or (bacterio)pheophytin (Type II RC). Notably, the reduced cofactor must be located at a relatively long distance (~ 20 Å) from the oxidized special pair to diminish the probability of charge recombination between them. Following the reduction by P*, A- is re-oxidized by a quinone molecule (Q) [Moser et al., 1992; Moser et al., 1993]. In Type I reaction centers, Q is phylloquinone, whereas in Type II reaction centers, it is plastoquinone in higher organisms (plants, algae) or ubi/menaquinone in others, such as purple photosynthetic bacteria for example. The electron transfer From P* to Q takes place within the range of picoseconds [Moser et al., 2003].
From the following step of the reductive electron flow, the Type I and Type II pathways are essentially different in the terms of events that lead to the ejection of the reducing equivalents from the RC. In Type I RC the electron is passed to the series of three iron-sulfur (4Fe4S) clusters, which are located within the RC (as shown in Figure 1), and then to ferredoxin, small soluble protein that aims the reducing equivalent to different pathways. In Type II RC the electron is delivered to a tightly bound quinone QA followed by a mobile quinone QB. When QB receives a second electron from the next photochemical turnover, it is protonated to form a quinol, which diffuses away from the RC into the lipid matrix, as described in the section.
Less sophisticated organisms such as heliobacteria and purple photosynthetic bacteria posses either Type I or Type II RC. In O2 evolving organisms, such as cyanobacteria, algae and higher plants, Type I (PSI) and Type II (PSII) RCs are coupled and act in series to drive oxygenic photosynthesis (Figure 1) as was firstly shown by M. Avron (1971) and earlier proposed by R. Hill and F. Bendall (1960). Despite the fact that PSI and PSII have derived from the symbiotic linkage of the Type I and Type II photosynthetic bacterium, they now differ from their ancestors in molecular detail [Mulkidjanian and Junge 1997; Mulkidjanian et al., 2003; Archer and Barber, 2004; Shi et al., 2005]. However, the fundamental principle that energy storage is accomplished by rapidly separating the initial oxidants and reductants of the primary charge separation to avoid wasteful recombination reactions, remained the major preserved motive throughout the evolution of RCs.
Overall, in oxygenic photosynthesis, excitation of PSI special pair generates a strong reductant, capable of reducing ferredoxin (followed by NADP+ reduction). Excitation of PSII special pair generates a strong oxidant capable of oxidizing water to oxygen. The overall electron transfer sequence is completed by the transfer of an electron between the reaction centers of PSII and PSI (Figure 1) and thus a sufficient energy for carbon fixation is provided. The functional redox coupling between the two RCs is accomplished by the cytochrome b 6 f (section I.III.I.).
An efficient light-harvesting step is critical for the success of photosynthesis and therefore photosynthetic organisms have advanced sophisticated pigment networks, called antenna systems [for reviews see Green and Parson, 2003]. Antenna systems have been evolved to engage the light harvesting into a broader light spectrum, increasing the effective absorption of RCs, which would be inefficient otherwise. The light, once captured by antenna is competently distributed, in the terms of excitation energy, to the RC for charge separation reaction and transmembrane electron transfer, presented in the previous section. This chain of events is achieved over a hierarchy of time scales and distances, with a remarkable efficiency [Sarovar et al., 2009; Bradler et al., 2009].
Antenna systems lack any photochemical activity of their own, and they only increase the number of pigments associated with each photochemical complex. The concept of antenna pigment system began to develop by R. Emerson and W. Arnold (1932), who showed that a cooperative cluster of chlorophyll molecules is required for releasing oxygen in light. Four years later H. Gaffron and K. Wohl explained these experiments by implying that most chlorophyll molecules are engaged in conveying the absorbed quanta, which may be efficiently transferred from one chlorophyll to another, and ultimately to the RC (originally referred as “photoenzyme ”). This provided an important ground for the following work by the Dutch biophysicist L. Duysens, which led him to conclude that chlorophylls excited by photons transfer the electronic excited state to neighboring compounds until it is trapped at the RC. This conception was entitled as "the true driving power of a photosynthetic unit" in Duysens’s famous Ph.D. dissertation (1952).
During the course of evolution, the natural selection have shaped and refined photosynthetic antenna systems according to unique ecological niches of corresponding organisms. Hence, antennas have adopted widely varied architectures in various classes of photosynthetic life forms (for structural reviews see Fromme, 2008). Despite the extraordinary diversity, all antenna systems share some basic characteristics and requirements that are critical for their functions and discussed below. A more detailed description of arrangements and compositions of specific antenna types relevant to this Ph.D. thesis are found in sections I.VI.IV. and I.VII.V. By incorporating many pigments into a single unit, antenna system maximizes the photosynthetic unit efficiency. In respect to the definition of efficiency, two factors have to be considered separately. First, the quantum efficiency q, which describes the number nRC of excitation quanta reaching the RC relative to the number nabs of quanta absorbed [Scheer, 2003]. Photosynthetic units are capable of reaching the quantum efficiency q of ~100%.
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There are many fundamental unsolved questions relating to the physical principles by which natural antenna systems retain such a high quantum yield, which is considerably unreachable by valid artificial photosynthetic systems [Noy et al., 2006; Amunts and Nelson, 2009b]. Thus, this is another aspect, which emphasizes the significance of characterization of the photosynthetic antennas’ spatial arrangement using X-ray crystallography method. The crystal structure information distinguished from other analytical methods by the sheer richness of the data that it provides: not only does it give the precise three-dimensional structure and geometry of individual molecules, but also vital information about how molecules interact with each other, which is essential starting point for understanding the design of natural light harvesting.
Second factor, the energetic efficiency e, which is defined by the energy eRC used by the RC for charge separation, relative to the energy eABS of a photon absorbed by the light-harvesting apparatus. Therefore, e relates to the energy gap between a lightquantum absorbed by the antenna and the excitation energy of the RC. In some photosynthetic units e can be quite low [Scheer, 2003].
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Functionally, for the light absorption and excitation energy transfer processes to be efficient, the overall transfer rate must operate on a time scale of femtoseconds [Sener et al., 2004], which is faster than the singlet lifetimes of chlorophyll (see below) [Duysens, 1964]. This is also the range at which the transfer time between neighboring pigment molecules in antenna systems has been estimated [Gobets and van Grondelle, 2001]. The rate of energy transfer in antenna systems depends mainly on the overlap of the donor-acceptor fluorescence emission spectrum. In other words, the energy lost as the donor decays to the ground state must match the energy gained as the acceptor excited. Resonance energy transfer represents a quantum mechanical resonance between two states with the same energy. For further physical mechanisms by which the energy transfer occurs, see Renger et al., 2001; Parson and Nagarajan, 2003; Scholes and Fleming, 2005; van Grondelle and Novoderezhkin, 2006; Adolphs and Renger, 2006; Engel et al., 2007; Renger, 2009; Cheng and Fleming, 2009; Collini et al., 2010.
Structurally, a cardinal requirement for energy transfer process to be efficient is that the location of each pair of energy donor-acceptor pigments would be within the range of ~16 Å. This center to center distance is favorable for delocalized electronically excited states [Forster, 1948; Forster, 1965]. This key criterion is accomplished by the protein scaffolding that enables strong couplings between appropriately oriented pigment molecules. It is noteworthy that based on structural information only, site energies and excitonic couplings of excitations between pigments can be determined using modern quantum chemical methods straightforwardly [Cheng and Fleming, 2009]. For an inter-pigment distance of > 10 Å, the excitonic coupling is determined by the Coulomb interaction between transitions [Alden et al., 1997; Cory et al., 1998; Hsu et al., 2008], which can be calculated using transition density cube method [Krueger et al., 1998; Scholes et al., 1999; Jordanides et al., 2001; Jordanides et al., 2004]. Therefore, elemental features of sunlight absorption by photosynthetic organisms may be precisely revealed by high resolution X-ray crystal structures coupled with modern calculations techniques, providing a powerful methodology.
For example, recently, Renger and coworkers calculated the site energies of seven bacteriochlorophylls in the Fenna-Matthews-Olson (FMO) complex from green sulfur bacteria using available structural information [Muh et al., 2007]. Importantly, this study suggested new energy-tuning mechanisms, which have not been considered previously. However, we do not know if these mechanisms are universal, and thus detailed modeling of other light-harvesting complexes would be highly informative in this context.
Another critical but less well-explored aspect of the energy transfer efficiency is the influence of protein environment [reviewed by Scholes 2003; Cheng and Fleming, 2009]. Interactions between pigments and their protein environment are among major contributors to the spectral properties and energy transfer characteristics. Protein moiety is able to modify excited states to alter their transition dipole densities. It was also shown that polarization of protein alters to some extent chlorophyll coupling, which is strictly dependant on relative orientation and position of donor/acceptor sites [Hsu et al., 2001]. In addition, hydrogen bonding between chlorin ring and H-atoms of amino acid residues was shown to contribute a positive shift of 60-100mV per bond to the potential of the electrostatic influence of positively charged residues in the immediate neighborhood of a chlorophyll [Lin et al., 1994; Mulkidjanian, 1999]. For a Schiff-base chlorophyll it was shown that its protonation increases the redox potential by ~310mV in vitro [Hanson et al., 1984; Maggiora et al., 1985].
Thus, evolution has realized the tuning of excitation energy transfer not only by clearly visible major structural features, but also through minor but important structural changes within the chlorophyll microenvironment. Taking this into consideration more sophisticated and realistic quantum mechanical models can be developed to calculate excitonic couplings within protein environments [Iozzi et al., 2004; Curutchet and Mennucci, 2005]. One such attempt was performed recently using time-dependent density functional theory, which considered a simple electrostatic model that includes the whole protein in atomic detail [Adolphs et al., 2008].
Overall, the accurate description of specific interactions between pigment molecules and protein residues is crucial for understanding of electronic excitations in photosynthetic antenna systems. However, since chlorophylls cannot be fully reconstituted within a protein moiety by available in vitro methods, the precise protein environment of pigment molecules cannot be observable other than by X-ray crystallographic studies aiming intact photosynthetic complexes. In fact, the available high resolution crystal structures of major photosynthetic complexes have already revolutionized our understanding of photosynthetic light harvesting by providing direct observations of antennas' engineering principles [Jordan et al., 2001; Camara- Artigas et al., 2003; Liu et al., 2004]. Further structural investigation is highly desired.
We will now analyze the antenna systems in more details by examining their functional building blocks - the light-harvesting pigments.
The two main pigment categories in the antenna systems of oxygenic photosynthesis, responsible for the absorption and tunneling of the excitation energy, are chlorophylls and carotenoids. Some of the key features of these pigments are described below.
Chlorophyll is a primary light-harvesting pigment in oxygenic photosynthesis. It was isolated for the first time by two French chemists, J. Pelletier and J. Caventou (1817), and was shown to be involved in oxygen evolution by the German botanist J. Sachs (1864). Chlorophyll's main absorption bands are in the red (~660nm, low energy) and blue (~450nm, high energy) regions of the visible spectrum, but they hardly absorb green-yellow light (~530nm, intermediate energy).
Chlorophyll consists of a tetrapyrrole ring (four pyrrol rings interconnected through one-carbon bridge in a cyclic fashion with overall size of ~1x1 nm) that binds magnesium atom (Mg) in the center (Figure 2). In addition to the four pyrrol ring, chlorophylls further contain a fifth isocyclic, oxygen containing ring, which able to produce hydrogen bonds either directly with amino acids or through water molecules buried in the protein (see example in section I.VI.III.II.). In photosynthetic units, chlorophylls are usually bound to specific protein pockets by direct ligation of the central Mg and/or via oxygen atoms. The long phytol chain attached to the periphery of the ring provides hydrophobicity that also aids binding to proteins. This hydrophobicity significantly influences the conformation of the tetrapyrrole ring (without interfering with the first solvation shell of the chromophore) and the removal of the phytol chain was shown to increase the ring flexibility [Fiedor et al., 2008].
Abbildung in dieser Leseprobe nicht enthalten
Figure 2 - Chemical structures of photosynthetic pigments.
Chlorophyll on the left and carotene on the right panel.
The central magnesium ion is not directly involved in the photosynthetic function and has no effect on the solvatochromism of chlorophyll -electron system. Rather, the chelation of Mg has a flattening effect on the tetrapyrrole ring [Fiedor et al., 2008], which absorbs the photon due to its well adapted electronic structure. The extensive usage of hemes in photosynthetic organisms as both, the electron carriers (cytochrome b 6 f, Alric et al., 2005) and photoinactivation protectors (Cyt b 559 in PSII, Morais et al., 2001), raises the interesting question - why hemes do not form the basis of any known antenna system instead of chlorophylls? A possible explanation is of two folds [Green et al., 2003]: 1) An antenna system built with hemes would have a less suitable absorption spectrum; 2) The excited states of hemes decay on picoseconds time scales by internal processes involving the iron atom. An excited heme thus has a little opportunity to transfer its energy to another molecule, and an excitation in a cluster of hemes would be degraded rapidly to heat. Replacing Fe by Mg extends the lifetime of the excited state up to nanoseconds range, allowing the energy transfer process to compete easily with the singlet lifetime. Therefore, chlorophylls, but not hemes, are to form complicated and yet precisely arranged antenna systems, which are designed to absorb photons and efficiently convey the excitation energy downhill to the special pair in the heart of RCs.
Carotenoids form another important group of photosynthetic pigments, which partially fill the chlorophyll absorption gap. They consist of a polyene chain of alternating single and double bonds, terminated by two rings. Carotenoids can differ in length, ring type and isomeric form, which also tune their absorption spectra. The most common type in photosynthetic units is called carotene, which consists only of carbon and hydrogen (Figure 2). The spectroscopic properties of carotenes are quite different from those of chlorophylls. Carotenes in the ground state (S0) absorb blue light reaching the singlet excited state (S2). The first excited state (S1) can not be populated from the ground state by photons absorption due to symmetry reasons. Carotenes are associated with pigment-protein complexes mainly involving hydrophobic interactions [Gilmore and Yamamoto, 1991; Giuffra et al., 1996; Croce et al., 2002a; Paulsen et al., 2003; Castelletti et al., 2003; Croce et al., 2004]. The association with proteins strongly red-shift absorption spectra of the carotenes, compared to one in the organic solvents. This shift represents a lowering of the S2 transition level, due to the mutual polarisability of carotenes and the protein environment [Plumley and Schmidt, 1987; Schmid et al., 2002].
Importantly, carotene’s excited-state energy can be transferred to chlorophyll, thereby enlarging light-harvesting capacity. Some close contacts between the two types of pigments are clearly seen in several sufficiently resolved crystal structures of photosynthetic complexes [Jordan et al., 2001; Liu et al., 2004; Guskov et al., 2009; Amunts et al., 2010]. In addition, carotenoids serve a significant role in photoprotection against light induced damage, in the case of over-excitation of the photosynthetic system [Yamamoto and Bassi, 1996; Frank and Cogdell, 1996]. The three main features that enable carotenoids to implement this role are: relative flexibility, rather high density of vibration states (which together result in very rapid internal conversion) and low energy of carotenoid triplets (which is below that of chlorophylls and that of singlet oxygen) [Scheer, 2003]. These key features, supplemented by the strongly forbidden S0-S1 excitation, enable carotenoids to act either as filters that reduce the amount of blue and near UV-light [Ben-Amotz et al., 1989] or as quenchers of 1) chlorophyll excited singlet states [Crimi et al., 2001]; 2) chlorophyll triplets [Hoff, 1993; Volk et al., 1993; Law and Cogdell, 1998]; 3) reactive oxygen species [Fiedor et al., 1993; Edge and Truscott, 1999; Young and Lowe, 2001; El-Agamey et al., 2004; Fiedor et al., 2005]. It is important to note that detailed quenching mechanisms are still a matter of debate [Mozzo et al., 2008; Berera et al., 2009; Triantaphylidès and Havaux, 2009; Bode et al., 2009; de Bianchi, 2010; Gruszeki, 2010].
Overall, different spectral properties of various light-harvesting pigments (and therefore their ability to perform efficient photoprotection) are complemented by vital structural support provided mainly by the protein moiety. As it was already mentioned, protein structures not only define pigment spatial arrangement and their proper orientation but also have a large effect on pigment spectroscopic properties. Taken together, these allow photosynthetic organisms to absorb and effectively consume the sunlight at wide spectrum of wavelengths available at the Earth's surface.
Since R. Hill and F. Bendall postulated the existence of the two light-driven steps, this hypothesis represented the cage of a skyscraper that is still nearing completion. First experimental proof for the functional performance of the two photosystems was provided in series of publications in 1961 by several laboratories [Duysens et al., 1961; Witt et al., 1961a, b Kok and Hoch, 1961, Kok, 1961], which independently demonstrated that an intermediate reaction component can be oxidized by far red light, that is, by PSI, and can be reversed to the reduced state only by shorter wavelength light < 700 nm, that is, by PSII. The concept was advanced later by L. Duysens and J. Amesz (1962) / L. Duysens and H. Sweers (1963), who sanctioned in definitive way that the photochemical reactions accomplished into two distinct photosystems working in series. In their study they reported results obtained from the red algae Porphyridium cruentum, in which two different pigment systems, absorbing light at two slightly different wavelengths, were clearly identified: one produced the oxidation of water (PSII), other the reduction of NADP+ (PSI).
Since these discoveries, the genuinely aspired challenge was to address central questions pertaining to the mechanism and evolution of the photosystems. To deal with the arduous tasks, a great number of scientists entered the scene and tackled the subject at biophysical, biochemical and structural levels [reviewed by Witt, 2004; Fromme and Mathis, 2004; Govindjee and Krogmann, 2004; Barber, 2004; Pennazio, 2008]. Essentially, the basic functional elements of the two photosystems and their component electron donors and acceptors were revealed by spectroscopic methods in the 1960s. The organization of these constituents within the photosynthetic membrane was elucidated by electrochromic band shift analysis. With the development of structural biology, a direct examination of the three dimensional structures became possible. Hence, the ultimate question for the last 20 years turned out to be: how the biological significance of photosystems is matched by their structural elements. However, structure determination is a complex process, and being such, it deserves a particular attention. Therefore, before depicting structural basis of PSI and PSII sunlight conversion process, we shall briefly address issues relevant to this Ph.D. thesis concerning the methodology, particularly the aspects and rationales of protein structural characterization.
The main methods for the three dimensional description of biomacromolecules are X- ray crystallography, nuclear magnetic resonance (NMR), and electron microscopy (EM). Among the three, the currently most powerful and predominately used technique aiming protein structures (~ 90% of total structures solved) is the X-ray crystallography [Worldwide Protein Data Bank, http://www.wwpdb.org]. An X-ray crystal structure provides overview of macroscopic protein architecture, which enable of granting direct crucial insights into mechanism of action to such extent that is not reachable by any other of the available techniques. Thus, some of the principal questions cannot be answered otherwise but throughout revealing a detailed crystal structure. Since, structures of individual components of large complexes often provide little insight into the mechanism of the assembly of which they are a part, it is essential to focus efforts on undertaking crystallographic studies of the whole and intact photosynthetic complexes.
However, PSI and PSII coordinate numerous pigments and comprise at least 19 protein subunits each, presenting a formidable challenge for the structural analysis [reviewed by Iwata and Barber, 2004; Vacha et al., 2005; Muh et al., 2008; Amunts and Nelson, 2009a]. In addition, PSI and PSII are complexes of membrane proteins, a class of proteins for which structural information is scarce. Whereas more than 70000 structures of soluble proteins have been determined [RSCB Protein Data Bank, http://www.rcsb.org], less than 250 different membrane protein structures have been unraveled so far, and almost all of these are from bacterial origin [Baker, 2010; Membrane Protein Data Bank, http://www.mpdb.ul.ie]. The reason for this is our limited ability to manipulate proteins bearing hydrophobic/amphiphilic surfaces that are usually enveloped with membrane lipid. Essential aspects of handling membrane proteins, while keeping them both active and suitable for crystallization, are discussed below.
Protein crystallography has been progressed significantly since the time of the giant pioneers such as M. Perutz, J. Kendrew, D. Phillips and D. Blow, however the fundamental methodology remained the same and it is composed of the four major steps: 1) obtaining a sufficient amount of pure and homogeneous protein; 2) formation of high quality crystals; 3) measuring and collecting the diffraction data from crystals; 4) diffraction data analysis and the following structure determination. The aspects of both usefulness and difficulties of this process are well documented in some of the recent reviews [Ilari and Savino, 2008; Wlodawer et al., 2008; Adams et al., 2009; DeLucas and DeLucas, 2009] with special emphasis on large macromolecular complexes [Mueller et al., 2007], membrane proteins [Xiong et al., 2007; Carpenter et al., 2008; Caffrey, 2009; Blois and Bowie, 2009; Sonoda et al., 2010] and photosynthetic pigment-protein structures [Fromme and Grotjohann, 2009; Allen et al., 2009; Li and Chang, 2009; Barros and Kühlbrandt, 2009]. Here, we specify only the key points, which are important for further discussion.
1) Pure and homogenous solution, in which all molecules are as identical as possible, is an essential prerequisite for protein crystallization. Previously, we reported that the quality of plant PSI crystals is highly related to the purity of preparation and even a very small contamination by ATP synthase and/or PSII prevents the crystal formation of PSI [Amunts et al., 2005]. Earlier experiments with cyanobacterial PSI crystallization showed that the presence of one monomer per 10,000 trimeric molecules prevents the growth of large well-ordered single crystals of trimeric PSI [Fromme and Witt, 1998; Fromme, 2003]. Further improvements of methods for the isolation of trimeric PSI from monomeric PSI, solely led to the improvement of resolution from 6 Å to 4 Å resolution. Therefore, not only that it is crucial to achieve a highest possible purification degree, but also to maximally reduce heterogeneity and remove impaired and partly denatured PSI complexes prior crystallization trials.
2) The stage of the formation of well diffracting crystals (highly ordered arrangement of protein molecules) is the bottleneck of crystal structure determination process, and therefore deserves a more detailed description. Despite recent technical and methodological advances, it still remains a major obstacle for X-ray crystallography. Protein crystals are composed of molecules that are precisely arranged in three- dimensional arrays. The smallest building block of the crystal is known as the unit cell, and the crystal can be considered to be composed of unit cells stacked next to each other. As originally suggested in the 1800s by M. Bravais, the arrangement of unit cells can be classified, according to the three lengths and angles associated with each unit cell and their symmetry relationship, into different space groups [International Tables for Crystallography, http://it.iucr.org].
The crystallization process can be divided into two major steps. First, nucleation, where crystalline nucleus is formed by ordered aggregation of protein molecules. This initial event may, as a result of statistical fluctuations, be the coalescence of enough molecules into an aggregate of sufficient dimensions. Essentially, crystallization from solution, like the formation of ice from water, represents a phase change. Upon reaching a critical size, defined by the competition of the ratio of the surface area of the aggregate to its volume [Feher and Kam, 1985; Boistelle and Astier, 1988], an organized protein crystalline aggregate creates a stable and viable nucleus, capable of further growth. From this point energy barrier has been surmounted and the phase transition leads to a more energetically favorable state of the system. Which means that inter-protein interactions with the nucleus become energetically feasible and the following stage begins, crystal growth. Numerous protein molecules aggregate with the existing nucleus in an ordered fashion faster than others separate out [for kinetic and thermodynamic parameters that characterize this process see Chernov et al., 1997; Drenth and Haas, 1992]. The association is accomplished through relatively weak, non-covalent bonds, hydrophobic interactions and salt bridges involving carboxylate anions and cationic amino or guanidinium groups immersed in a water environment [McPherson, 1999]. The energetic gain of aggregating to developing organized crystal increases as the joining monomers enable to generate more interactions with its neighbors. In this manner, the crystal spontaneously develops, as long as the solution is saturated. The success of crystallization depends on finding appropriate reagents and conditions (precipitants, buffers, different additives, temperature and other various parameters) under which protein molecules are optimally arranged in an ordered fashion while protein concentration is held in the supersaturated state, so that the crystals can separate out.
The major obstacle with macromolecular crystallization is that crystal formation and growth constantly competes with protein amorphous precipitate. The energetic barrier of amorphous sediment creation is much lower, which make this state kinetically favored. In other words, small amorphous aggregate is more stable than small crystal, despite the energetic preference of mature crystal upon amorphous sediment. Therefore, precipitates tend to appear first in the equilibration process and often preclude crystal formation. The situation is particularly acute because the competition is promoted by high levels of supersaturation. Thus, the objective of a crystallographer is to find ways of inducing crystal formation (and growth) while discouraging precipitation.
However, it is worth to note, that there have been numerous reports in the literature of the transformation observed between amorphous precipitates and crystalline state [McPherson, 1999]. When this occurs, the precipitate is usually observed to form almost immediately after the crystallization trial is composed (due to its kinetic favorability), however, because crystal lattice represents a lower energy potential, it is eventually ultimately favored. It is therefore, the energy difference that drives the conversion from an amorphous state to an ordered one [Ng et al., 1996].
There are additional origins of crystal disorder such as thermal motions, solvent molecules collisions, etc., which are valid also after the completion of crystal formation. Obviously, these disorders disrupt the quality of structural data produced from crystals, and eventually weaken detail separation ability (resolution) of the crystal structure. The effect of these processes is much more radical when it concerns large protein molecules such as photosystems.
3) Once crystals have been obtained, diffraction measurements are performed utilizing undulator radiation at intense X-ray beams available at synchrotron sources. The crystals are aligned and exposed to the X-ray beam with the diffraction measurement being detected in real time by an electronic detector. In order to determine the crystal structure, it is necessary to measure the intensity of every X-ray diffraction spot derived from the protein crystal. Since the data are recorded electronically, the measurements can be performed relatively quickly. The space group can be identified based on the arrangement of the diffraction peaks in the resulting image, while the intensity is related to the composition of the unit cell.
However, one of the major, and especially relevant to the structural biology of photosystems, obstacles in the effective measurement and data collection at synchrotrons is the radiation damage suffered by protein crystals [Garman and Owen, 2005; Garman and Nave, 2009; Holton, 2009]. With the advent of third-generation synchrotrons and the provision of undulator fed beamlines, the observation of radiation damage has become commonplace, manifested as a reduction in diffracting power and a loss of high-resolution reflections even at cryotemperatures of ~100 K [Owen et al., 2006; Massover, 2007]. Generally, radiation damage is an undesirable component of the experiment and can result in erroneous structural detail in the final model [Nave and Garman, 2006; Weiss et al., 2005]. Therefore, the characterization of radiation damage has recently became an important developing area for structural biologists [Burmeister, 2000; Ravelli and McSweeney, 2000; Ravelli et al., 2002; Weik et al., 2000, Weik et al., 2002; Leiros et al., 2006]. Incident X-rays can induce specific structural changes in a well defined sequence, with some amino acids being more susceptible than others. Specific structural damage to covalent bonds occurs first to disulfide bridges, glutamates and aspartates are then decarboxylated, tyrosine residues lose their hydroxyl group, and subsequently the C-S bonds in methionines are cleaved. Thus, special care is required when interpreting structures, which may have been modified by X-ray damage during the data collection. Metalloproteins are particularly susceptible to partial reduction during the diffraction experiment, and may not be in their native state by the end of the data collection [Carugo and Djinovic Carugo, 2005]. Thus in the context of structural biology of photosystems radiation damage has now became an issue of wide concern [Barber and Murray, 2008].
4) The diffraction can be considered to arise from the summation of vectors, called structure factors, which have both an amplitude and phase, but the above described measurement of the intensity provides only the amplitude. Formally, in order to determine the structure, both are required in a Fourier series involving the summation of the structure factors . The summation yields density of electrons at a given point in space, since it is the electrons that scatter the X-rays. For this reason, the outcome of the data analysis is an electron density map into which a structural model, consisting of amino acid residues, must fit.
Four experimental approaches can be used to provide the missing information concerning the phases: multiple anomalous dispersion (MAD), which involves measurements at several wavelengths around the transition energy for a metal bound to the protein [Hendrickson, 1999]; single anomalous dispersion (SAD), which uses a single dataset at a single appropriate wavelength, is technically simpler than MAD but requires accurate estimation of anomalous intensity differences [Hendrickson and Teeter, 1981; Wang, 1985]; multiple isomorphous replacement (MIR), where the protein is modified such that a metal is incorporated and the diffraction is compared to the protein without the metal, with the measurements all being at a single wavelength [Hendrickson and Ogata, 1997; Bella and Rossmann, 1998]; molecular replacement (MR), if an existing structure that is highly homologous to the unknown structure is available [Rossmann, 1972].
These approaches provide the means for generating the phases for the diffraction data and allowing the electron density to be calculated. Fitting of the maps involves identifying the atoms that give rise to each region of the density. While this is largely manually done on a graphics terminal by a crystallographer, increasingly, the analysis of electron density maps is being performed directly by computer programs. In our case, phases were determined using the molecular replacement method. To eliminate model bias, a composite omit map was calculated in the final stages of the refinement. The phase determination from the model was accurate enough to generate an electron density map that revealed additional features that were missing from the original model.
Because of their extremely hydrophobic character, membrane proteins are only stable in solution with appropriate detergents are added [reviewed by Seddon et al., 2004; Linke, 2009]. Detergents mimic the lipid bilayer by covering the hydrophobic surface of membrane proteins with their alkyl chains. In this manner membrane proteins are prevented from generating non-specific hydrophobic interactions, minimizing the denaturation and precipitation processes [le Maire et al., 2000; Gohon and Popot, 2003; Arnold and Linke, 2008]. However, the influence from detergents has two impeding characteristics not present in soluble protein crystals. The first is that protein-protein contacts, which are essential for the formation of the crystal lattice, can only be mediated through the hydrophilic surfaces of membrane proteins extending out of the detergent micelles [Michel and Oesterhelt, 1980; Garavito and Rosenbusch, 1980; Deisenhofer and Michel, 1989]. A method, which expand hydrophilic surface of proteins using antibody fragments, has been developed [Michel, 1991]. In theory, increasing the hydrophilic surface facilitates the formation of crystals, however in practice, it had only a limited success and has not became a commonly used strategy for acquiring membrane protein crystals.
The second impediment for crystallization stems from the fact that detergent solubilized membrane proteins suffer from lipid depletion following their purification. In other words, detergent micelles usually cause spatial- and time-dependent disorder [Zulauf, 1991; Nollert et al., 2002]. Lose of lipids affects stabilization in the way, which may be destructive to membrane proteins [Garavito and Ferguson-Miller, 2001; Gouaux and White, 2001]. Thus, the problem of membrane protein purification may not center on obtaining the highest possible purity, but rather an optimum purity that does not cause depletion of the native lipid content [Zhang et al., 2003]. In this respect, two roles of lipids have been discussed in the literature: (i) a functional role that was inferred from structure studies on bovine cytochrome c oxidase [Mizushima et al., 1999]; bacteriorhodopsin [Essen et al., 1998], yeast cyt bc 1 complex [Lange et al., 2001], E. coli outer membrane protein OmpX [Fernandez et al., 2002] and RC of Rb. Sphaeroides [McAuley et al., 1999; Camara-Artigas et al., 2001; Camara-Artigas et al., 2002]; (ii) a structural role has been proposed for the specific lipids digalactosyl-diacylglycerol and phosphatidylglycerol in crystallization of the plant light-harvesting complex II [Liu et al., 2004; Standfuss et al., 2005], and for cardiolipin bound to the RC of Rb. Sphaeroides where it forms a partial peripheral shell around the hydrophobic protein core, acting as boundary lipid at the protein- detergent interface [Camara-Artigas et al., 2002]. Biophysical studies suggest that in the presence of detergent and absence of essential lipids, the much smaller lateral pressure exerted by the detergent micelle may result in greater conformational freedom of the side chains and backbone of the protein [Cantor, 1999; PebayPeyroula and Rosenbusch, 2001]. Supplemented by an increased H2O accessibility and resulting lability [Landau and Rosenbusch, 1996], crystallization of membrane proteins may be frustrated. Therefore, addition of a small amount of pure lipids to the protein-detergent complex may be considered [Zhang et al., 2003].
Detergent effectiveness varies between different proteins, in addition micelles, which optimally fitted for solubilization, do not necessary fit into the crystal lattice of the protein. Hence, a given detergent may be suitable for solubilizing and stabilizing a specific membrane protein, but may hinder the protein molecule interactions that are necessary for crystallization to occur [Kühlbrandt, 1988]. If this is the case, changing or adding other kinds of detergent is necessary for the purposes of crystallization. Choice of detergent is, therefore, the most important factor in membrane protein crystallization, apart from the stability and homogeneity of the protein target [Lin and Guidotti, 2009]. In some cases, mixed detergents are required when the desired results cannot be obtained with a single kind of detergent. For example, well formed crystals of the oxygen-evolving PSII reaction centers from spinach and pea could only be obtained by utilizing detergent mixtures [Adir, 1999].
Intensive biochemical work and systematic protein characterization, in combination with comprehensive screening for the optimally suitable detergents, is probably the most efficient strategy to cope with the difficulties of membrane protein crystallization [Hunte and Michel, 2002]. S. Iwata summarized crystallization conditions that have been used successfully for crystallizing of -helical membrane proteins, and designed a special screening kit for membrane proteins, which is widely used for confining initial conditions for membrane proteins crystallization [Newstead, et al., 2008]. However, finding an initial crystallization condition for membrane proteins to yield small crystals is not the major hurdle and is only one step for the structural biologist. In most cases, the initial crystallization conditions, which yield small crystals, are not the optimal conditions for crystallization, and hence we need to adjust various parameters in order to obtain the best quality crystals. This further optimization to generate crystals that are suitable for X-ray crystallography is probably the most challenging task on the road towards a successful structure determination.
A milestone of extensive efforts to elucidate photosynthetic complexes structures was the determination of the purple bacterial RC crystal structure at 2.9 Å by H. Michel, J. Deisenhofer and R. Huber (1985). This significant achievement represented a turning point in membrane protein crystallization history and spurred many structural biologists to pursue the structures of large membrane proteins. The two later major breakthroughs in unraveling photosystems were the elucidations of the 2.5 Å resolution X-ray crystal structure of the trimeric PSI from cyanobacteria TS elongates [Jordan et al., 2001] and the 3.5. Å resolution structure of dimeric PSII from the same organism [Ferreira et al., 2004]. These and recently improved structural data [Besiadka et al., 2004; Loll et al., 2005; Guskov et al., 2009], provide now the tools to extend the structural comparison to the complete structures of PSI and PSII, revealing amazing similarity in their principal elements.
First of all, the structures have confirmed the idea of a modular architecture of all photosystems and photosynthetic reaction centers [Blankenship, 1992; Schubert et al., 1998; Rhee et al., 1998; Baymann et al., 2001; reviewed by Grotjohann et al., 2004; Hohmann-Marriott and Blankenship, 2008; Krauss, 2008] The photosystems and all the known reaction centers contain a core structure of pseudo- C2 symmetry, with a central domain consisting of 2 x 5 transmembrane -helices that bind the intramembrane cofactors of the electron transport chain. In PSII, this central domain consists of the similarly organized subunits D1 and D2, while in PSI it is the C- terminal parts of the homologous proteins, PsaA and PsaB. An additional conserved structural motif of 2 x 6 transmembrane -helices is formed by the core antenna proteins CP43 and CP47 in PSII, and the N-terminal components of PsaA/PsaB in PSI, arranged as trimers of dimers. Therefore, in PSII the reaction center core (2 x 5) and the core antenna (2 x 6) are formed by distinct protein subunits, whereas in PSI the hetero-dimer PsaA/PsaB constitutes both domains. The overall structure formed by the 2 x 5, plus the 2 x 6 transmembrane -helices also obeys the pseudo C2 symmetry.
From the evolutionary point of view, it is especially remarkable that the most pronounced differences between the transmembrane -helices of both photosystems exist in those helices that coordinate most of the electron transport chain components. In both RCs, these -helices (namely "D" in PSII and "10" in PSI) form the center of hemi-spheroid structures, and coordinate the primary electron donor (P680 in PSII and P700 in PSI) by a pair of histidines. However, there is no further sequence homology between those helices. In addition, their length and tilt is also significantly different. While -helices "10" of PSI form a V-shaped arrangement, which ends at a distance of about 8 Å from the stromal membrane surface and extends into the lumen, providing part of the binding site for plastocyanine/cytochrome c 6, in PSII the two "D" -helices form a much longer X-like structure extending into the stroma. Based on the fact that the most important -helices with respect to the coordination of the electron transfer chain cofactors differ much more than the helices in the antenna region, Grotjohann et al., (2004) suggested that strong evolutionary pressure led to the divergent evolution of both photosystems, which in turn was a pre-requisite for the development of efficient oxygenic photosynthesis.
We begin the comprehensive description of the two photosystems from PSII, because this is the primary complex that drives photosynthesis and consequently provides electrons to PSI. Understanding PSII on the structural level is highly important for this Ph.D. thesis, as it is vastly related to PSI. Functionally, the two complexes are synchronized and in respect to a whole plant leaf scale, one cannot fully operate without the other. Therefore, PSII analysis will serve us for developing a more fruitful discussion on PSI at a later stage. However, due to the limit of space, the obtained results will be described here only briefly and usually presented in the form of conclusions rather than highlighting the methodologies and rationales supporting them.
PSII is unique as it is the only enzyme on Earth, which is able to split water into protons and molecular oxygen by use of visible light. In view of its importance, the structure and function of PSII has been studied extensively in the past several decades. In general, PSII contains a type-II RC that catalyzes light-driven electron transport from water to a plastoquinone. The heart of PSII is known to be largely conserved from cyanobacteria to plants. From a crystallographic point of view, it took about one decade from the first single crystals of PSII capable of light-induced oxygen production [Zouni et al., 1998; Zouni et al., 2000] to the most recent crystal structure refined to 2.9 Å resolution [Guskov et al., 2009]. Between 2001 and 2005, several crystallographic models with resolutions ranging from 3.8 to 3.0 Å have been reported by different groups [Zouni et al., 2001; Fromme et al., 2002; Kamiya et al., 2003; Besiadka et al., 2004; Ferreira et al., 2004; Loll et al., 2005]. These were determined from two different but closely related thermophilic bacteria: TS elongates and TS vulcanus. Recently, the 3D-crystallization of PSII from red algae has been reported as well [Adachi et al., 2009], however PSII structure from plants has not been determined by X-ray crystallography yet. Therefore, further discussion of PSII structure is based on the available crystallographic models for bacteria, whereas features of unique plant structural elements (presented in section I.VI.IV) are based mostly on spectroscopic data and recent EM analysis [Caffari et al., 2009].
All available crystallographic models of PSII determined the architecture of functional homodimeric complex with a molecular weight of about 750 kDa (upon finishing writing these lines a 3.6 Å resolution crystal structure of monomeric PSII from TS elongates was submitted by A. Zouni group). However, it is also important to note that very recently it was suggested that PSII complexes exist in the form of a monomer in vivo and the two distinct monomers become a dimer during the purification steps incorporating detergents between their interfaces [Takahashi et al., 2009].
The crystallized PSII homodimer consists of 20 different protein subunits [Guskov et al., 2009] Of these, 17 span the membrane (PsbA to PsbF, PsbH to PsbM, PsbT, PsbX to PsbZ and ycf12) and three are membrane-extrinsic (PsbO, PsbU and PsbV), as shown in Figure 3. Each monomer of PSII harbors nearly 100 cofactors, in reasonable agreement with a chemical analysis of cofactor composition [Kern et al., 2005]. According to the 2.9 Å resolution structure [Guskov et al., 2009], PSII has the following cofactor inventory per monomer: 35 chlorophyll a (Chl a), two pheophytin a (Pheo a), two heme (cyt b 559 and cyt c 550), 12 ß-carotene, three plastoquinones (QA, QB, and QC), 25 integral lipid, and seven dodecyl maltoside (DDM) molecules, one calcium (Ca2 +) and four manganese ions of the Mn4Ca cluster, one chloride ion close to this cluster, two additional Ca2 + ions, and one non-heme iron (Fe2 +) with an associated bicarbonate ion, summing up to 96 cofactors. The complexity of this molecular framework mirrors a diversity of the function carried out by PSII as highlighted in recent review papers focused on various performing aspects of this intricate machine [Renger and Renger, 2008; Muh et al., 2008; Semenov et al., 2008; Ivanov et al., 2008; Barber, 2008; Barber, 2009; Pospisil, 2009; Guskov et al., 2010].
Abbildung in dieser Leseprobe nicht enthalten
Figure 3 - General architecture of PSII.
On the left: view of the PSII homodimer along the membrane plane (cytoplasm, top; lumen, bottom). The 25 lipid and 7 detergent molecules per monomer are shown in space- filling mode (carbon, yellow; oxygen, red), protein subunits in gray, and the three membrane-extrinsic subunits in green (PsbO), violet (PsbU) and blue (PsbV). On the right: view of PSII from the cytoplasmic side (membrane-extrinsic subunits omitted). The monomer-monomer interface is indicated by a black dashed line. Helical parts are shown as cylinders, and the subunits PsbA (D1, yellow), PsbB (CP47, red), PsbC (CP43, magenta), PsbD (D2, orange), cyt b 559 (cyan, subunits PsbE, , and PsbF, ) and the remaining eleven small subunits (light blue) are labeled in the left monomer. Cofactors are shown in stick mode: Chl (green), Car (orange), heme (blue). In right monomer, all protein subunits in gray, lipids and detergents shown as spheres (carbon, yellow; oxygen, red).
The two large subunits PsbA and PsbD, traditionally referred to as D1 and D2, respectively, form the central part of the membrane-intrinsic part of PSII and harbor the RC, where light-induced electron transfer is initiated. The RC is flanked by the two large subunits PsbB (CP47) and PsbC (CP43), which bind 16 and 13 Chl a, respectively, and constitute the core antenna system of PSII. Besides their function as antenna proteins, PsbB and PsbC also play an important role in the stabilization of the core complex and the Mn4Ca cluster (see below) by virtue of their large membrane- extrinsic loops extending into the lumen and being in contact with the extrinsic proteins.
The other 13 membrane-intrinsic protein subunits of PSII are of a relatively low molecular weight and feature only one transmembrane -helix with the exception of PsbZ, which shows two (Figure 3). Until now PsbY could only be modeled as poly- alanine due to poor electron density in this particular area, most probably because this subunit is loosely attached to the complex [Kashino et al., 2007]. PsbX is located next to PsbH and PsbD. Subunit Ycf12 is situated next to subunits PsbJ, PsbK and PsbZ.
The two monomers of the PSII homodimer are related by a non-crystallographic C2 symmetry. At the monomer-monomer interface, the small subunits PsbL, PsbM, and PsbT form a three-helix bundle (Figure 3). According to the 2.9 Å resolution model [Guskov et al., 2009], the two symmetry-related subunits PsbM and PsbM’ interact by virtue of a heptad motif of aliphatic side chains as in a leucine zipper. Similar motifs have been shown to promote the assembly of transmembrane protein segments [Gurezka et al., 1999]. However, deleting PsbM in a mutant of the mesophilic cyanobacterium Synechocystis sp. PCC 6803 is not sufficient to prevent dimer formation, but the additional deletion of PsbT is required [Bentley et al., 2008; Watanabe et al., 2009]. As shown in Figure 3, the distance between the two PsbT subunits in the homodimer seems to be rather large, but they are in contact with integral lipids that are located at the monomer-monomer interface. Therefore, an important role in dimer formation was ascribed to both lipids and small protein subunits [Guskov et al., 2010].
PSII is known to bind a number of membrane-extrinsic subunits of largely unknown function at its lumenal side. The identity of these subunits varies between different types of organisms [Roose et al., 2007; Enami et al., 2008]. The crystallized PSII from TS elongatus contains PsbO, PsbU, and PsbV. Whereas PsbU binds no cofactors, PsbV is a heme protein (cyt c 550), and PsbO harbors a Ca2 + binding site. The role of these subunits is mainly in the formation of educt and product channels associated with the water oxidation process [Ishikita et al., 2006; Murray and Barber, 2007; Ho, 2008; Ho and Styring, 2008; Guskov et al., 2009; Gabdulkhakov et al., 2009]. It is possible that more extrinsic subunits exist, which are lost during purification procedures or bind temporarily to PSII [Guskov et al., 2010]. In red algae, for example, PsbQ is present and was suggested to affect the crystal packing of dimeric PSII [Adachi et al., 2009]. Green algae and plants possess neither PsbU nor PsbV, but PsbQ, PsbP and PsbR instead [Roose et al., 2007; Enami et al., 2008]. PsbQ of spinach and PsbP of tobacco could be separately crystallized, and the structures were determined to 1.49 Å [Calderone et al., 2003; Balsera et al., 2005] and 1.6 Å [Ifuku et al., 2004] respectively.
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