University of Groningen Structures of Photosynthetic Membrane

University of Groningen

Structures of photosynthetic membrane complexes Semchonok, Dmitry Alexandrovich

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Introduction

Photosynthesis is the processes whereby plants, algae and some other groups of organisms convert light into chemically fixed energy. The topic of this thesis deals with the primary steps in photosynthesis, which takes place in specialized photosynthetic membranes. Complex proteins inside photosynthetic membranes catalyse the primary steps. A number of such proteins were structurally characterised by electron microscopy. In this introduction chapter, we will discuss some basic aspects of both photosynthesis and electron microscopy analysis. Further, the proteins and the corresponding organisms that have been studied will be introduced.

Some basic facts about photosynthesis Photosynthesis is the first main issue to be introduced. It is the process whereby green plants, algae, cyanobacteria, photosynthetic bacteria and certain other organisms transform light energy into chemical energy that can be later released to fuel the organisms' activities: namely to convert water, carbon dioxide and minerals into oxygen and energy-rich organic compounds in a direct or indirect way. The main organic compounds are carbohydrate molecules, sugars, lipids and proteins. Besides being vital for the life of the photosynthetic organisms, they serve as food for all other living creatures (Encyclopedia Britannica, 2010). Thus the meaning of the word “photosynthesis” – from the Greek φῶς, phōs, "light", and σύνθεσις, synthesis, "putting together" (Ke, 2001) has significance in the most broadest sense. Looking around and watching to fossils in old stones it seems that photosynthesis exists forever. However, how old is photosynthesis more precisely? It is generally believed the earth was formed around 4.54 billion years ago by accretion from the solar nebula and that life on earth began between 3.8 – 3.5 billion years ago (Noffke et al., 2013). In the Archean Era (3.9–2.5 billion years ago) the earliest photosynthetic activity was carried out by bacteria that did not evolve oxygen. These so-called anoxygenic photosynthetic bacteria are assumed to have used reductants such as H2, H2S, or ferrous iron, but not H2O. Oxygenic photosynthesis carried out by 6

cyanobacteria is thought to have been developed later (Olson, 2006). Evidences indicate that cyanobacteria evolved about 2 billion years ago leading to oxygen accumulation in the atmosphere. The presence of oxygen and the production of foodstuffs by higher plants made the existence of heterotrophs such as humans possible. Presently, the total biomass produced annually by plant photosynthesis amounts to about two hundred billion tons (Ke, 2001). Living creatures, including humans, consume the photosynthesised foodstuffs and gain energy from them by “respiration”, a process by which the organic compounds are oxidized back to carbon dioxide and water. Photosynthesis therefore serves as a vital link between the light energy of the sun and all living creatures (Ke, 2001).

History of the study of photosynthesis Photosynthesis, as one of the most important processes on Earth, takes a central position in plant cell science. The first known scientific view on the process of photosynthesis was expressed by Aristotle (384 – 322 BC), the “father of biology”. He compared the soil (earth) to the stomach and assumed earth as the stomach of plants, as soon as they gain their nutrients directly from earth and water without having a “proper” digestive system. After observations of Aristotle, there is a big gap in the history of the research for 2000 years until the 17th century when the wave of interest to photosynthesis and plant science research arose with a new force. The next scientist who focused on photosynthesis was Jan Baptist van Helmont (1579 – 1644), an early modern period Flemish chemist, physiologist, and physician, who considered water to be the source of life and the basic nutrient for plants. Therefore, he devised an experiment by which he showed that small potted willows could thrive on soil and water alone while they gain their substance (weight) solely from the “water” as the weight of the soil in the pots did not decrease significantly. His works were collected and edited by his son Franciscus Mercurius van Helmont and the book “Ortus medicinae, vel opera et opuscula omnia” was published by Lodewijk Elzevir in Amsterdam in 1648 (NOYES, 1895) wherein the term “gas” (from the Greek word chaos) was used for the first time. After that time, the study of photosynthesis was slowly increasing. In our review, we cannot omit Marcello Malpighi (1628 – 1694), an Italian physician and biologist regarded as one of the fathers of microscopical anatomy and histology, who studied the anatomy of plants and insects concisely by making use of the

microscope. He claimed that plants take up nutrients which are dissolved in water via their roots. Overall, a pleiad of eminent scientists for over 300 years has made an invaluable contribution to the study of photosynthesis. Among them are persons as Robert Boyle, Joseph Priestley, Jan Ingenhousz, Julius Sachs, Kliment Arkadievich Timiryazev, Albert Einstein and many others. Between 1925–53, a number of novel techniques and methods enabled much more detailed research in plant physiology. It was the period that controlled growth in climatized growth chambers, ultracentrifugation, electron microscopy, x-ray-diffraction, thin layer and gas liquid chromatography and fluorescence spectrophotometry became available. Most of these techniques are still prevailing in modern research. Many of the central concepts of photosynthesis were established around the middle of the 20th century and at the same time, its basic mechanisms were clarified in more detail. For example, measurements of photosynthetic efficiency (quantum yield) at different wavelengths of light (Emerson and Lewis, 1943) led to the insight that two distinct forms of chlorophyll (Chl) must be excited in oxygenic photosynthesis. These results suggested the concept of two photochemical systems. The reaction centre pigments of photosystem II (PSII) and photosystem I (PSI) (P680 and P700, respectively) were found by studying changes in light absorbance in the red region (Kok, 1959), (Döring et al., 1969), (Tanaka and Makino, 2009). Chls with absorbance maxima corresponding to these specific wavelengths were proposed as the final light sink. These Chls were shown to drive electron transfer by charge separation. The linkage of electron transfer and CO2 assimilation was suggested by studies on Hill oxidant (Hill, 1937). A linear electron transport system with two light-driven reactions (Z scheme) was proposed based upon observations of the redox state of cytochromes (Hill and Bendall, 1960), (Duysens et al., 1961) and photophosphorylation was found to be associated with thylakoid fragments (Arnon et al., 1954). The metabolic pathway that assimilates carbon by fixation of CO2 was 14 discovered by Melvin Calvin's group, who used CO2 radioactive tracers in the 1950s (Bassham et al., 1950). This was the first significant discovery in biochemistry made using radioactive tracers, for which Calvin received the chemistry Noble prize in 1961. The primary reaction of CO2 fixation is catalyzed by ribulose-1,5-bisphosphate carboxylase/oxygenase, commonly known by the abbreviation Rubisco (Weissbach et al., 1956), initially called Fraction 1 protein (Wildman and Bonner, 1947). Rubisco is the most abundant protein in the world, largely because it is also the most inefficient

–1 one with the lowest catalytic turnover rate (1–3 s ). Another CO2 fixation pathway was found later in sugarcane (Hartt and Kortschak, 1964), (Hatch and Slack, 1966). It was named C4 photosynthesis (Tanaka and Makino, 2009) to discriminate it from the much more common C3 type of photosynthesis. One of the next breakthroughs to our understanding of photosynthesis was achieved by Hartmut Michel and Johann Deisenhofer. They made crystals of the photosynthetic reaction centre from Rhodopseudomonas viridis, an anaerobic photosynthetic bacterium, and used x-ray crystallography to determine its three-dimensional structure (Deisenhofer and Michel, 1989). In 1988, they shared the Nobel Prize in Chemistry together with Robert Huber for this groundbreaking research.

Present situation Current research on photosynthesis covers a wide range of aspects, from basic science to multiple types applications. To start with science, we have to consider physical, biochemical, physiological and ecological aspects. The central concepts of photosynthesis have been elucidated, including the primary steps of light capturing by PSI and PSII and electron and proton flow (“light reactions”). We also know the components involved in metabolic pathway of sugar synthesis in the Calvin cycle and other pathways (“dark reactions”). Parts of these processes can be understood at the molecular level, because high-resolution protein structures are available. Yet, there are still numerous open questions, concerning, for instance, the regulation of PSII under excess of light. There is no detailed model how plant PSII is modified under non-photochemical quenching. Photosynthesis is part of a number of diverse physiological processes that need to stay in harmonic balance. One of the primary functions of it is to control the redox state of cells, by changing the enzyme activity and many other cellular processes (Buchanan and Balmer, 2005), (Hisabori et al. 2007). Here is still a lot to discover. One regulatory function of photosynthesis is carried out by causing the generation of reactive oxygen species, that now is appreciated as being a regulatory factor for many biological processes, instead of being only a harmful side product of photosynthesis (Wagner et al., 2004), (Beck, 2005). Chlorophyll molecules also play a dual function. Chlorophylls are a major component of light- harvesting in photosynthesis. Precursor molecules of chlorophyll, however, can act as a chloroplast-derived signal and are involved in regulating the cell cycle (Kobayashi et al., 2009). In lightweight of these new facts, it appears

necessary to re-evaluate the function(s), both potential and demonstrated, of photosynthesis from a range of perspectives. There are also new developments in ecology, although a closer discussion is outside the scope of this thesis. But meanwhile, chlorophyll fluorescence and gas exchange measurements, developed especially for photosynthesis research, are now widely used in stress biology and ecology (Tanaka and Makino, 2009). Use of photosynthetic research can help to comprehend the ecological phenomena and even the global environments (Farquhar et al., 1980), (De Pury and Farquhar, 1997), (Monsi et al., 2005). Nowadays the process of photosynthesis is included as integral part of many simulations of the future programs. Concerning application, one field that is actively developing is artificial photosynthesis. One line of research is to reconstruct the process of photosynthesis or a part of it with the help of artificially made proteins. Recently, researchers have constructed a molecular catalyser - mononuclear ruthenium complex, that can oxidize water to oxygen very rapidly, comparable to that of photosystem II (Duan et al., 2012). As a result, it become possible to reach speeds of catalytic activity approximately to those of natural photosynthesis - about 100 to 400 turnovers per seconds (Kaftan, 1999). Scientists have now reached over 300 turnovers per seconds with their artificial photosynthesis models. The research findings play a critical role for the future use of solar energy and other renewable energy sources. Improving of the efficiency of photosynthesis in plants, algae or cyanobacteria is another wide field of work. The light-harvesting within the photosystems and antenna complexes is hard to improve, but a key for the improvement lies in modifications of proteins regulating photosynthesis. Plants easily put their photosynthesis capacity on hold under high light or stress conditions to prevent photodamage, at the cost of optimal speed of growth or productivity. Modern biotechnology have made it possible to manipulate photosynthesis using molecular genetic technology as well (Andrews and Whitney, 2003), (Raines, 2006). Modification of the proteins that involved in regulation processes may be a key to speed up photosynthesis. This could lead to positive influences on crop productivity, as photosynthetic rates have frequently been correlated with biomass accretion (Kruger and Volin, 2006). It looks obvious that future research will open the new horizons for research in this direction.

About EM Microscopy is a scientific discipline that implies the use of microscopes to view samples and objects that cannot be seen within the resolution range of the normal eye. Knowledge about the structure of different objects leads to better understanding of the world around us. Biology in particular relies heavily on microscopy to gather information, and this scientific tool is in daily use all over the world (“What is Microscopy?” n.d.). There are several well-known branches of microscopy: optical, electron, and scanning probe microscopy. Optical microscopy, which involves the use of visible light, was the first form to be introduced. It is also known as “light microscopy.” Having obvious limits connected with the wavelength of visible light this type of microscopy gave way to other one - electron microscopy (EM) - that was invented in a 20th century. In EM, the object is illuminated with an electron beam. Electron microscopy produces excellent detail, but the equipment is costly and the specimens must be prepared very precisely in order to get useful results.

Resolution in Microscopy A microscope is a perfect tool to enhance the visibility of small details of your specimen. At the same time, this enhancement is subject to some physical laws that need some explanation to understand its possibilities and limits. Resolution (or resolving power) is defined as the closest spacing of two points which can be resolved by the microscope as separate entities. In simple words, it is the possibility to see two spots separately. As soon as an aperture of any physical lens has fixed parameters, a point source of light that is going through is not seen as a point of light but as the diffraction pattern of the instrument aperture. That diffraction pattern is called the Airy disk. Based on the Rayleigh criterion that was formulated in 1896 the formula for resolution is the following:

1,22λ R = ; 훮훢 푐표푛푑푒푛푠푒푟+푁퐴 표푏푗푒푐푡푖푣푒 where R = resolution, measured as distance, depends on the angular aperture α; NA = numerical aperture, equal to ηsin(훼/2), which depends on the diameter of the lens and its focal length, η = refractive index of the medium between the lens and specimen and λ = the wavelength of light illuminating/emanating from the sample.

Practically the angle α/2 cannot exceed 70°. The maximum value NA of a condenser or objective lens in air is 0.95. The NA of a light microscope lens can be increased to typically 1.45 by an immersion lens is, using immersion oil. The shortest wavelength of visible light is 400 nm. Then it follows that the resolution can be: 1.22×400 nm R = = 203 nm; 1.45+ 0.95

For many years it was thought that the resolution of about 200 nm was a hard limit for light microscopy. However, resolution below this theoretical limit can be achieved using near-field scanning optical microscope or a diffraction technique called 4Pi STED microscopy (Pohl et al., 1984). But generally speaking, the resolution is practically limited by the wavelength of light you are using. The wavelength of the electrons depends on accelerating voltage and in a 10 kV scanning electron microscope is then 12.2 x 10−12 m (12.2 pm), while in a 200 kV transmission electron microscope (TEM) the wavelength is 2.5 pm. This implies that the theoretical resolution limit for an electron microscope is 4 orders of magnitude higher than for a light microscope. Based on that idea the electron microscope, where the source of illumination is the electron beam, has a real advantage for nanostructural analysis.

Electron microscopy The Transmission Electron Microscope was the first type of Electron Microscope to be developed and its basic configuration is similar to that of the Light Transmission Microscope, except that a focused beam of electrons is used instead of visible light to "see through" the specimen. The first electron microscope was developed by Max Knoll and Ernst Ruska in Germany in 1931 (Ruska, 1986). In general the procedure performed during EM analysis includes the following steps: a beam of electrons is formed in high vacuum (by an electron gun). Electrons are accelerated towards the specimen, while the beam is narrowed and focused using metal apertures and magnetic lenses of the condenser system into a thin, focused, monochromatic beam. Then the sample is irradiated by the beam causing the interactions inside the sample that affect the electron beam. These interactions and effects are detected and transformed into an image. One problem on the way to get a high resolution image is the poor quality of electron lenses. This causes the presence of different lens aberrations: spherical, chromatic, astigmatism. In order to optimise the imaging 12

properties different approaches are taken. The first is based on limitation of the beam with small apertures to reduce the effect of spherical aberration. The weak side of it comes from the resolution limit formula where the resolution has linear dependence on the size of numerical aperture. A more modern approach is using different correctors and filters. With aberration correction all electrons are focused within the region of interest. As an example, to correct the current spherical aberration in electron microscopes is to introduce a corrector that produces negative spherical aberration. The combination of negative and positive aberration of the objective lens gives a total of zero spherical aberration.

Contrast The most important aspect of electron microscopy is the presence of contrast. Leaving the detailed theory of contrast formation for what it’s worth, we will briefly summarize the basic ideas of it. “Contrast” is the appearance of an object feature in an image. In Electron microscopy, the contrast for simplicity can be decomposed into several components: scattering and phase contrast.

Scattering contrast The electrons that hit the specimen in an electron microscope are partly scattered by interaction with atoms in the specimen. The interaction of primary electrons with nuclei is generally elastic - the energy of the primary electron (and the related wavelength of the electron wave) almost does not change. However, a phase shift of the electron wave will generally occur. Interaction of the primary electrons with electrons in the specimen does lead to a significant loss of energy for the primary electrons and is therefore inelastic. The objective aperture stops electrons scattered (elastically) over relatively large angles. The presence of local differences in scattering power in the specimen leads thus to contrast in the electron microscopic image. This form of contrast is called amplitude or scattering contrast.

Phase contrast The phase shift of the electron wave due to elastic interactions by itself does not lead to contrast in the image. However, lens aberrations (particularly spherical aberration, Cs) and defocusing (Δf) introduce an additional phase shift, which is a function of the spatial frequency (ν = 1/d) in the back-focal plane: 4 4 2 2 χ(ν) = 2π/λ (-Cs(λ ν /4) + Δf(λ ν /2) ) 13

The contrast in the image, which is caused by interference of the scattered waves (which have the additional phase shift) with the unscattered wave (which has no additional shift), is called phase contrast. The contrast transfer function (CTF) for phase contrast is given by Sin(χ(ν)):

4 4 2 2 Sin(χ(ν)) = Sin[2π/λ (-Cs(λ ν /4) + Δf(λ ν /2) )], where: Cs (the quality of objective lens defined by spherical aberration coefficient), λ (wave-length defined by accelerating voltage), ∆f (the defocus value), ν (spatial frequency)

Considering the importance of contrast in microscopy, there have been many attempts to improve it. With regard to electron microscopy the implementation of this technique requires certain conditions with respect to the sample preparation. For example, the necessity to work with biological samples under vacuum using an electron beam that causes radiation damage, has led to three major sample preparation techniques: negative staining, plastic embedding and cryo-technique. In negative staining the specimen is embedded in a heavy-metal salt. As a result the background becomes stained and sample stays untouched, but dehydrated. In the opposite case the method is called positive staining. Practically speaking, water around the sample is substituted by heavy metal salt that also protects it from radiation damage. This is an easy, rapid, qualitative method for examining the structure of isolated organelles, individual macromolecules and viruses at the EM level. The first paper that described the negative staining technique was published by Sidney Brenner and Robert Horne in 1959 (Brenner and Horne, 1959). The usefulness of the method was first reported by Hugh Huxley at a conference in Stockholm in 1956. But the very first one to apply it was Cecil Hall in 1955, before Hugh Huxley, who had used the negative staining method without pointing out its value, and Friedrich Krause, who had seen a similar outlining phenomenon already in 1937 (Maunsbach and Afzelius, 1998). Many stains are used for negative staining, of which ammonium molybdate, uranyl acetate, uranyl formate, uranyl oxalate, phosphotungstic acid, sodium silicotungstate, methylamine tungstate and auroglucothionate have shown the best results. These solutions have been chosen for their ability to dry down in a glassy state and scatter electrons well, leaving the matter itself

relatively intact. The structures which can be negatively stained reveal details which are much smaller than those obtained with any type of light microscopy. But talking about the advantages of this technique we also have to mention some downfalls (Table 1). One of them is a limit in maximum achievable resolution that for most specimens equals ~1.5 nm, even when the studied object has smaller details, which are typical for proteins and many other types of macromolecules. This limit is probably set by the “grain size” of the stain.

Table 1. Advantages and disadvantages of the negative staining technique

Advantages Disadvantages good signal-to-noise ratio for small molecules are prone to structural molecules collapse simple to apply high background from surrounding stain resistant to radiation possible chemical interactions works well on heterogeneous preps distortions due to ionic strength and low pH can induce preferred orientation limited resolution 3-D reconstruction is possible imaging under non-native conditions

Biological electron microscopy obtained new life with the invention of a cryo-technique by Dubochet and McDowall in 1981 in order to preserve the specimen (Dubochet and McDowall, 1981). The overall idea of the technique is to rapidly freeze the biological sample at liquid nitrogen temperature ~ 90 K in a thin layer of ice. The biological sample for the moment of freezing stays in water or buffer solution. If the freezing step is done fast enough, the liquid water transforms to an amorphous state of ice, without the formation of crystals. Therefore, the biological sample stays almost intact in the solution. Table 2 describes some advantages and disadvantages of this technique.

Table 2. Advantages and disadvantages of cryo-technique

Advantages Disadvantages no artefacts due to fixation, low signal-to-noise-ratio dehydration or staining preservation of native sensitive to radiation conformation random orientation technically challenging good contrast at high defocus difficult to distinguish between different orientations vs. conformations higher resolution info than freezing artefacts negative stain

The images that can be obtained by cryo/negative staining techniques are two–dimensional (2D) projections owing to the large depth of focus of the TEM. Cryo-EM, together with computer reconstruction techniques, open the possibility for making a 3D image of the specimen structure using large amounts of image projections with different tilts. There are three general techniques for achieving 3D imaging: single-particle electron microscopy (cryo-EM), cryo-electron tomography (cryo-ET) and cryo-electron crystallography (cryo-EC). These methods are based on the fact that the parallel projection of a 3D specimen is equivalent to a slice in the 3D Fourier space of the object. To form the total 3D reconstruction it is necessary to obtain different slices in Fourier space. In tomography the specimen is rotated in small increments in the microscope and a (weighted) back-projection method is typically used to form the 3D reconstruction (Beck et al., 2004), (Masters and Vermaas, 2001). Alternatively, for single- particle analysis, identical copies of the specimen occur in many different orientations, and these images are used to reconstruct the 3D structure. The technique of cryo-EC can be used to determine the 3D structures of macromolecular assemblies from 2D crystals (Masters et al., 2001). Again, tilting of the specimen in the microscope is necessary to obtain 3D information.

Photosynthesis Phototrophic organisms use light for their energy. In the broadest sense, there are two types of photosynthesis. The result of photosynthesis is common for both of them – the light (photons) is used to synthesise and store chemical energy in the form of ATP and / or NADPH. Phototrophy can be divided in chlorophyll-based chlorophototrophy that as it follows from the name has chlorophyll in the centre of the process and rhodopsin- based retinalophototrophy. Both ways of light harvesting are very different. Retinalophototrophy is very simple. In the model organism, Halobacterium halobium, there is just one protein responsible for light-harvesting and conversion, which is bacteriorhodopsin (Haupts et al., 1999). It has only one retinal pigment molecule attached. In contrast, protein molecules functioning in chlorophototrophic organisms have multiple copies of interconnected chlorophyll molecules attached. Bacterial reaction centres with a peripheral antenna system and plant photosystems are large protein complexes which contain dozens of chlorophyll copies. A calculation presented by Boekema et al., 2013 indicates a pigment density per membrane surface of 1 retinal per 11.1 nm2, which is about 4 times lower than for the bacteriochlorophylls of purple bacteria. This means in practice that retinalophototrophic organisms are restricted to places with high light levels, such as the Great Salt Lake and the Dead Sea. Chlorotrophic organisms such as green photosynthetic bacteria can, on the other hand, even grow over 1000 meter deep in the ocean where light is extremely limited (Boekema et al., 2013). Therefore in practice only chlorophototrophy is associated with photosynthesis. It can be divided into oxygenic and non-oxygenic photosynthesis and its most relevant aspects in the context of this thesis are discussed in the next parts. Oxygenic photosynthesis produces most of the organic matter on Earth, as well as almost all of its oxygen. The primary steps in this process – converting light energy to usable chemical energy – are carried out by four multisubunit membrane–protein complexes. Two of the complexes, PSI and PSII, work as molecular photovoltaics by emitting electrons upon the absorbance of light energy. The third complex, cytochrome b6f (cyt. b6f), mediates the transport of electrons between the two PSs and further contributes to the formation of a proton-motive force by pumping protons over the membrane (Nevo et al., 2012). The last complex, an F-type ATPase (CF1CF0-ATP synthase), converts the proton motive force into ready-to-use ATP molecules by rotary catalysis. Supplementing these complexes are two small mobile carriers which mediate the transport of electrons. A quinone

molecule, called plastoquinone (PQ), serves in plants the transport of electrons from PSII to cyt. b6f. The role of PQ is similar to ubiquinone, which is a quinone functioning in mitochondrial electron transport. A small, water-soluble copper-binding protein, called plastocyanin (PC) mediates electrons from the cyt. b6f complex to PSI (Nevo et al., 2012). In plants and algae, photosynthesis occurs in a special organelle, the chloroplast. An average leaf cell of higher plants may have about 40 – 50 chloroplasts. The size of chloroplast of land plants is 5–8 μm in diameter and 1–3 μm thick (Wise, 2006). They have an oval shape with a compartment structure inside, enclosed by three membrane systems: an outer and inner membrane and a thylakoid system. Inside the inner membrane is the stroma, a gel-like medium that fills most of the chloroplast including the thylakoids, which are disk-like stacks of folded but interconnected membranes. Thylakoids stacks, or grana stacks, surround the thylakoid lumen. Grana stacks are connected by stroma thylakoid membranes into the functional compartment. The nature of the biomolecules involved in this process long time remained a mystery (Rabinowitch, 1956). Some light was shed on the process in the middle of the last century, when the concept of two photosystems was introduced and confirmed by experimental evidence. The idea of System I and System II as physical parts of the photosynthetic apparatus and the seats of light-induced charge separation, each with specific antenna pigments, was first introduced by Duysens in 1960 (Duysens et al., 1961), (Duysens, 1989). Now we know that PSI and PSII with their antenna system, as well as cytochrome b6f complex, are exclusively located in the thylakoid membrane of oxygenic photosynthetic organism (Ort and Yocum, 1996), (Wydrzynski and Satoh, 2006), (Golbeck, 2006). The most familiar form of photosynthesis is non-cyclic photophosphorylation. It consists of two sets of pigments to excite in the order of PSII and PSI. Reaction center of PSI is better excited by light at about 700 nm, and is thus sometimes called P700. PSII cannot use photons of wavelength longer than 680 nm, and is thus sometimes called P-680. If both systems would be close together in the membrane, the lower-energy absorbing PSI would take most of the excitations of antenna proteins of PSII. Thus, there is a need for spatial separation. Hence the primary processes of photosynthesis in plants are located in different parts of the thylakoid membrane (Fromme and Mathis, 2004). PSI is exclusively present in the stroma thylakoid membranes whereas most of the PSII is present in the grana membranes.

Photosystem II Photosystem II (or water-plastoquinone oxidoreductase) is the first photosynthetic pigment protein complex in the light-dependent reactions of oxygenic photosynthesis. It is a homodimeric multisubunit protein–cofactor complex localised in the thylakoid membrane of plants, algae, and cyanobacteria. The major function of photosystem II is to transfer captured energy of light to the electron transport chain, reduction of plastoquinone, creation of a proton gradient across the thylakoid membrane and the oxidation of water which leads to the evolution of oxygen (Govindjee, 2006). The ability to produce oxygen is unique in nature, because there are no other proteins that can split water and this has millions of years ago crucially changed the conditions and living life on the Earth. The general equation of photosynthesis is the following:

+ + 2H2O+ 2PQ + 4H → O2 +4H + 2PQH2, where PQ stands for oxidized plastoquinone, and PQH2 for fully reduced plastoquinol.

The Photosystem II complex consists of multiple proteins and pigment molecules. Up to date it includes 33 subunits (Table 3). The core part from both cyanobacteria and plants contains well over 20 subunits. The large subunits PsabA-PsbD are common to both plants and cyanobacteria. PsbC (CP43) and PsbD (CP47) are the two largest subunits and form an internal antenna in the core. The names were derived from apparent masses of 43 and 47 kDa, respectively on gels. PsbA (D1) and PsbD (D2) form inside the core complex the photochemical reaction centre in which the charge separation and primary electron transfer reactions take place. They bind a special pair of chlorophyll molecules P680, which donates an electron to the electron transport system. In PSII, the electron is then passed to a pheophytin molecule, then to plastoquinone Qa and after to plastoquinone Qb. These PQ molecules are embedded in the D2 and D1 proteins. After obtaining two electrons the fully reduced Qb carries them to the cytochrome b6f complex. Lost electrons are transferred back from water to the P680 molecules that have delivered their electrons in the process. This reaction is + called water-oxidation and has the following equation: 2H2O → O2 + 4H + 4e−. The D1 and D2 proteins are by function analogous to the L and M proteins in the purple bacterial reaction centre (discussed below) and show

weak but definite sequence homology to the L and M subunits (Michel and Deisenhofer, 1988). Plants and cyanobacteria have slight differences in the small subunits with single transmembrane α-helices, but also have many components that are highly conserved. Among the conserved subunits, present in all PSII complexes, are the low molecular weight subunits PsbE and PsbF, that bind the high potential heme of cytochrome b559 (Cyt b559). The PSII core complex has several hydrophilic extrinsic proteins attached to its lumenal surface, which form a protein shield over the catalytic site of water splitting (Barber, 2014). Together, these proteins are called the oxygen evolving complex (OEC). Some of these proteins are common to all oxygenic photosynthetic organisms, others vary between different types of organisms. The OEC in higher plants and green alga includes: PsbO (33 kDa), PsbP (23 kDa), PsbQ (17 kDa) and PsbR (10 kDa) (Allahverdiyeva et al., 2013), while in cyanobacteria the five extrinsic proteins in OEC are: PsbO, PsbP- like, PsbQ-like and (Cyt) c550 (PsbV, 17 kDa) and PsbU (12 kDa (Bricker et al., 2012), (Govindjee, 2011). The total molecular mass of the monomeric PSII core complex is 350 kDa, which is slightly higher than the PSI core (see below). Although monomeric PSII subcomplexes are present in native thylakoid membranes (Danielsson et al., 2004), the PSII core complex normally exists as a dimer. The most recent and detailed published crystal structure of dimeric PSII core complex has a resolution of 1.9 Å from Thermosynechococcus vulcanus (Umena et al., 2011) (Fig. 1). It can be seen that the membrane-embedded part is composed of numerous transmembrane α-helices. One side of the complex (the stromal side at the top of model in frame A) is remarkably flat, whereas the other side shows the bulky oxygen-evolving complex. The four large subunits PsbA-PsbD have in total 22 transmembrane α-helices and are surrounded by a number of small membrane proteins (in green) which main function appears to keep the full complex in an active and stable conformation.

A B

Figure 1. Structure of Photosystem II from the thermophilic cyanobacterium Thermosynechococcus vulcanus at 1.9 Å resolution. View from the direction are A) parallel and B) perpendicular to the membrane plane. The protein subunits are colored individually: PsbA (D1) – blue, PsbD (D2) – cyan, PsbB (CP47) – hot pink, PsbC (CP43) – orange red. Source: PDB ID: 3ARC

There are no core complex structures of plant or algal PSII available. At the moment, the only available low-resolution structure of different PSII complex was obtained using electron microscopy techniques (Kouřil et al., 2012). The resolution of the obtained structure allows to analyse and understand the structure and partly the functioning of this complex, but only at the subunit level (Fig. 2). The model of Fig. 2 shows a central dimeric core complex (C). The monomeric structure of the cyanobacterial PSII has been fitted on the position of one monomer, although there may be small discrepancies between both systems. But it nevertheless shows how the plant core PSII is part of a larger particle, the PSII supercomplex. The light- harvesting components CP29 (Lhcb4), CP26 (Lhcb5), CP24 (Lhcb6) and the Light-Harvesting complex II (LHCII) are unique for plants and do not exist in cyanobacteria. In fact, there is not just one PSII supercomplex, because three different LHCII proteins, Lhcb1-3, are organized in heterotrimers. The standard large supercomplex of the model plant Arabidopsis thaliana binds 4 LHCII trimers. The innermost or S-trimers are connected by CP26 and CP29 to the core complex (Fig. 2). A more peripheral M-trimer is attached via CP29 and CP24. Because different combinations of trimers are possible, the supercomplexes have been named according to the presence of S and M trimers (Dekker and Boekema 2005). The particle presented in Fig. 2 contains two S trimers and two M trimers

and is named C2S2M2. Particles with one M trimer les (C2S2M) are common in spinach, together with C2S2 complexes. The latter are also dominant under high-light conditions, were the need of a large peripheral antenna is less important. The positions of the LHCII trimers are clearly definable within a map of negatively stained C2S2M2 supercomplexes (Caffarri et al., 2009). Although improving the resolution of these complexes in 2D still can shed some light on organisation them in vivo, getting 3D maps is the final goal. Some attempts in this direction have already been done (Nield et al., 2000), (Pagliano et al., 2013).

Figure 2. Structure of dimeric supercomplex of Photosystem II from higher plants, as determined by single particle electron microscopy. (A) The top view projection map of the complex, seen from the luminal side of the membrane. (B) The projection map overlaid with a dimer of two PII core complexes. The right side shows a detailed structure of the Photosystem II reaction centre and various light harvesting complexes, while the left side is labelled with C for reaction centre core, M and S for major light harvesting complexes LHCII and minor light harvesting complexes CP24, CP26, and CP29. The scale bar equals 10 nm. Source: (Kouřil et al., 2012).

Table 3. Subunit composition of plant and cyanobacterial Photosystem II

Subunit Gene Gene Mass Cofactors Function name locationa (kDa)b PSII-A (D1) psbA C 39 chlorophyll, Core reaction pheophytin, centre of quinone, Photosystem II 훽-carotene, Fe PSII-B psbB C 56 chlorophyll, Core antenna (CP47) 훽-carotene PSII-C psbC C 51 chlorophyll,훽- Core antenna (CP43) carotene PSII-D (D2) psbD C 39 chlorophyll, Core reaction pheophytin, centre of quinone, Photosystem II 훽-carotene, Fe PSII-E psbE C 9 heme Core reaction (cytb- centre of 559훼) Photosystem II PSII-F psbF C 4 heme Core reaction (cytb- centre of 559훽) Photosystem II PSII-H psbH C 8 phosphate Photoprotection,

QA to QB regulation PSII-I psbI C 4 Core reaction centre of Photosystem II PSII-J psbJ C 4 Assembly of Photosystem II PSII-K psbK C 4 Role in PSII assembly

PSII-L psbL C 4 Role in QA binding PSII-M psbM C 4 Role in PSII stability PSII-N psbN C 5 Role in PSII stability PSII-O psbO N 27 Stabilizes Mn- (OE33) cluster, Ca2+ and Cl− binding

PSII-P PsbP N 20 Ca2+ and Cl− binding (OE23)c PSII-Q PsbQ N 17 Ca2+ and Cl− binding (OE16)c PSII-Rc PsbR N 10 ? PSII-S PsbS N 22 chlorophyll, Antenna regulation (CP22) carotenoids by xanthophyll cycle PSII-T psbT C 3 Role in PSII stability (ycf8) d PSII-U psbU 14 Role in O2 evolution

PSII-V psbV 15 heme Role in O2 evolution (cytc-550)d PSII-Wc PsbW N 6 Role in PSII stability

PSII-X psbX C 4 Role in QA function PSII-Y PsbY N 4 Mn binding? (ycf32) PSII-Z psbZ C 9 Antenna-reaction (ycf9) centre interaction Psb27 slr1645 12 Role in PSII assembly Psb28 sll1398 13 Role in PSII assembly Psb29 sll1414 27 Role in PSII assembly Psb30 sll0047 3 Stabilize PSII (ycf12) e Psb31 12 Role in O2 evolution Psb32 sll1390 22 Role in PSII repair LHCII- Lhcb1 N 30 chlorophyll, Antenna function outerc carotenoids LHCII- Lhcb2 N 31 chlorophyll, Antenna function outerc carotenoids LHCIIa- Lhcb3 N 25 chlorophyll, Antenna function outerc carotenoids LHCII- Lhcb4 N 35 chlorophyll, Antenna function inner carotenoids (CP29)c

LHCII- Lhcb5 N 36 chlorophyll, Antenna function inner carotenoids (CP26)c

LHCII- Lhcb6 N 18 chlorophyll, Antenna function inner carotenoids (CP24)c aGene location applies only to eukaryotic organisms. C, chloroplast; N, nucleus bMass is actual mass based on gene sequence cFound only in eukaryotic organisms dFound only in cyanobacteria eFound only in marine diatoms (Blankenship, 2014)

Photosystem I Photosystem I (PSI) (or plastocyanin: ferredoxin oxidoreductase) is also a multisubunit pigment protein complex. It is the second photosystem in the photosynthetic light reactions of algae, plants, and some bacteria. Photosystem I is so named because it was discovered before photosystem II (Duysens, 1989) (Fig 3). PSI catalyses the light driven electron transfer from plastocyanin/cytochrome c6 on the lumenal side of the membrane to ferredoxin/flavodoxin at the stromal side by a chain of electron carriers (Fromme et al., 2001).

Prokaryotic and eukaryotic PSI structures In general, the minimal PSI complex consists of a monomeric core, with a substantial antenna. Photosystem I has approximately 100 chlorophyll molecules and 12-16 β-carotene molecules per monomeric core, which is larger than PSII. In the case of eukaryotic PSI there are some additional LHCI outer antenna subunits. The reaction centre of the photosystem I core is a heterodimeric protein core complex made by integral membrane proteins called PsaA and PsaB (82-83 kDa), that consists of 22 transmembrane helices and contains the special pair of dimeric chlorophylls P700. PSI evolved over 3.5 billion years ago from a simple homodimeric structure into a sophisticated apparatus that consists of a heterodimeric core. It harbours the intrinsic pigments and cofactors for excitation energy and electron transfer and a peripheral antenna providing additional pigments for efficient light-harvesting (Busch and Hippler, 2011). In comparison with

oxidizing photosystem II, photosystem I seems more reducing (Nelson and Yocum, 2006), (Golbeck, 2006). The PsaA/PsaB complex is surrounded by a variable amount of smaller proteins in the range of 4 to 25 kDa. Some of these proteins may serve as binding sites for the soluble electron carriers plastocyanin and ferredoxin. The functions of some other proteins are not well established (Table 4). One of the differences between prokaryotic cyanobacteria and eukaryotic organisms is the presence of some additional small proteins, such as PSI-H, PSI-G, PSI-N and PSI-K (Table 4). Some of these small subunits enable to adjust LHCI outer antenna complexes to the core. The plant-specific LHCI consists of different chlorophyll a/b binding proteins called Lhca’s, surrounding the core of PSI (Croce and Van Amerongen, 2011). The function of pigments is to harvest sunlight energy and to transfer the trapped excitation energy to the reaction centre (RC). Photosystem I of cyanobacteria exists in vivo in trimeric and monomeric form (Grotjohann and Fromme, 2005). A trimeric form of the PSI complex was firstly found in Synechococcus sp. by electron microscopy in 1987 (Boekema et al., 1987) and the detailed structure of the PSI trimer was determined using X-ray crystallography (Jordan et al., 2001) to 2.5 Å resolution, revealing the positions of the many cofactors. The PSI monomer of Synechococcus elongatus consists of 12 protein subunits, 96 chlorophyll a molecules, 22 carotenoids, three [4Fe4S] clusters and two phylloquinones. Furthermore, it has been discovered that four lipids are intrinsic components of PSI. The native trimer of PSI has a molecular mass of 1068 kDa and thereby it is the largest and most complex membrane protein that has been crystallized so far (Fromme et al., 2001). The trimer is, however, not the highest type of organization. Recently it was shown that PSI complex of some cyanobacteria is organized as a tetramer (Watanabe et al., 2014), (Li et al., 2014). A structural characterization indicated that the 4 monomers comprising this complex are arranged in a pseudotetrameric form, consisting of two dimers of dimers. After the cyanobactarial structure, the plant structure could also be resolved. The crystal structure of pea PSI and its refinements give a high impact to understanding of the structure and function of eukaryotic PSI (Amunts et al., 2007), (Ben-Shem et al., 2003), (Amunts et al., 2010), (Qin et al., 2015), (Mazor et al., 2015).

A B

C D

Figure 3. The Overall Structure of PSI of eukaryote (A,B) and prokaryote C,D,E. Each individual protein subunits is colored differently. Positions of several subunits are indicated. (A) View from the lumen. Chlorophylls and amino acids are shown in transparency. LHCI is comprised of Lhca1–4 chlorophyll binding proteins, assembled in a half-moon shape onto the RC. Binding of LHCI to RC is asymmetric, namely, much stronger on the PsaG pole than on the PsaK pole. PsaH prevents the trimerization of PSI. (B) View perpendicular to the membrane normal. Chlorophylls of LHCI are blue, gap chlorophylls are cyan, the rest of RC chlorophylls are green. The PsaC/D/E ridge forms a ferredoxin docking site. PsaF involved in the binding of plastocyanin. PsaN is found to interact with Lhca2/3 (Amunts and Nelson, 2009). Crystal structure of T. elongatus PSI monomer (PDB ID: 1JB0) shown from27 the lumen (C) and from the stromal side up (D). (E) Trimeric PSI complex constructed from 1JB0 structure shown luminal side up (PDB ID: 4FE1) Table 4. Subunit composition of Photosystem I

Subunit Gene Gene Mass Cofactors Function name name locationa (kDa)b chlorophyll, Core reaction PSI-A psaA C 84 quinone, centre of PSI, (PsaA) 훽-carotene, charge separation, Fe–S electron transport chlorophyll, Core reaction PSI-B psaB C 83 quinone, centre of PSI, (PsaB) 훽-carotene, charge separation, Fe–S electron transport PSI-C psaC C 9 Fe–S Role in electron

transport PSI-D PsaD N 18 Ferredoxin docking PSI-E PsaE N 10 Role in cyclic electron transport PSI-F PsaF N 17 Plastocyanin docking c PSI-G PsaG N 11 Role in QA binding PSI-Hc PsaH N 11 Interaction with LHCII PSI-I psaI C 4 ? PSI-J psaJ C 5 ? PSI-K PsaK N 9 Role in docking of LHCI PSI-L PsaL N 18 Trimer formation in cyanobacteria PSI-Md psaM 3 Trimer formation in cyanobacteria PSI-Nc PsaN N 10 Plastocyanin docking PSI-Xd psaX 3 chlorophyll Binding of IsiA antenna LHC-Ic Lhca1 N 22 chlorophyll, Antenna function (LHCI- carotenoid 720) LHC-Ic Lhca2 N 23 chlorophyll, Antenna function

(LHCI- carotenoid 680) LHC-Ic Lhca3 N 25 chlorophyll, Antenna function (LHCI- carotenoid 680) LHC-Ic Lhca4 N 21 chlorophyll, Antenna function (LHCI- carotenoid 720) aGene location applies only to eukaryotic organisms. C, chloroplast; N, nucleus bMass is actual mass based on gene sequence cFound only in eukaryotic organisms dFound only in cyanobacteria (Blankenship, 2014)

In plants, green and red algae as well as in diatoms PSI exists in monomeric form. Trimerization is prevented by the presence of the PSI-H (PsaH) subunit. The monomer consists of two functional parts: the core complex – reaction centre (RC) and the light-harvesting complex (LHCI). The core docks the components for the light-driven charge separation and the following electron transfer reactions (Busch and Hippler, 2011). Additionally, it binds approximately 100 chlorophylls (Chls) (Amunts et al., 2007) which serve as antenna system to collect light energy. This core antenna is extended by the light-harvesting complex (LHCI) which forms a crescent-shaped structure at the PsaF/PsaJ side of the core and is energetically coupled to the PSI core via the “gap chlorophylls” (Ben-Shem et al., 2003). Up to now 15 subunits (PsaA to L and PsaN to P) of the eukaryotic PSI core are known and the most recent refinement of the crystal structure of the PSI–LHCI complex to 3.3 Å identified in total 18 protein subunits, 173 Chls, 15 β-carotenes, 3 (4Fe–4S) clusters, and 2 phylloquinones (Amunts et al., 2010). Some protein subunits such as PsaG, PsaH, and PsaN are special to eukaryotes and are involved in binding the light harvesting polypeptides. The PSI core is highly conserved throughout evolution (Ben-Shem et al., 2003), while the LHCI complex shows a higher degree of variability in size, subunits composition and bound pigments, which is due to the large variety of different habitats photosynthetic organisms live in (Busch and Hippler, 2011). Especially in green algae, the LHCI antenna is substantially larger with 9 different copies of antenna subunits.

The detailed structure of PSI electron transfer cofactors and the pathway of electron transfer is illustrated in Fig.4. As in PSII in PSI the electron transport processes begin with transfer of the excitation energy from pigment to the reaction centre. Energy transfer delivers excitations to the core electron transfer cofactors (Melkozernov et al., 2006). Additional electron acceptors include a series of three membrane- associated iron–sulphur proteins, also known as Fe–S centres Fe–SX, Fe–SA, and Fe–SB. Fe–S centre Fe–SX is part of the P700-binding protein; centres Fe–SA and Fe–SB reside on PsaC, a protein of 8 kDa that is part of the PS-I reaction centre complex. Electrons are transferred through centres Fe–SA and Fe–SB to ferredoxin, a small, water-soluble iron–sulphur protein. The membrane-associated flavoprotein ferredoxin–NADP reductase (FNR) reduces NADP+ to NADPH, thus completing the sequence of noncyclic electron transport that begins with the oxidation of water (Karplus et al., 1991) (Fig. 4). In cyanobacteria ferredoxin is replaced by flavodoxin under iron-stress. Electron transport between PSII and PSI is mediated by plastohydroquinone, the cytochrome b6f complex and plastocyanin. The cytochrome b6f complex (see below) contains two β-type hemes and one c- type heme. The PS-I reaction centre appears to have some functional similarity to the reaction centre found in the anaerobic green sulphur bacteria and the heliobacteria. These bacteria contain low-potential Fe–S centres as early electron acceptors and are probably capable of ferredoxin-mediated NAD+ reduction similar to the NADP+ reduction function of photosystem I. There is almost certainly an evolutionary relationship between these complexes and photosystem I of oxygen-evolving organisms.

The structure and function of the cytochrome b6f complex Cytochromes are membrane-bound hemeproteins and are responsible for electron transport in a large number of organisms. They were fist described by MacMunn as myohematin or histohematin in 1884 (Munn, 1887). Cytochromes can exist in a monomeric form, such as the small cytochrome c protein and as a subunit of bigger complexes. Some of the larger cytochrome complexes are primarily for the generation of ATP via their proton pumping activity, which is coupled to electron transfer. There are several different types of cytochromes, and these are distinguished in a variety of ways, including by spectroscopy and sensitivity to specific

inhibitors. There are cytochromes a, b, and c — and each has further subtypes. In prokaryotic organisms, such as purple bacteria, the cytochrome bc1 complex is the main proton pump in the photosynthetic light reaction,

Figure 4. Structural model of the electron pathway for light induced electron transport from plastocyanin to ferredoxin in photosystem I. Chls (blue), quinones (black), the copper atom of plastocyanin (Pc) (blue), and Fe (red balls) and S (green balls) of the three iron-sulphur clusters and the Fd iron-sulphur clusters are depicted. Two tryptophan residues (light- blue and light-pink space-filling structures), implicated in electron transport from plastocyanin to P700, are also shown in the context of their secondary structural environment. (Nelson and Yocum, 2006).

whereas higher plants, cyanobacteria and algae have the b6f complex. These two cytochromes have similar function, although their subunit composition and structure differs. The cytochrome b6f complex is an essential player in noncyclic and cyclic electron flow (Baniulis et al., 2008), (Cramer et al., 2011), (Hasan et al., 2013). It occupies a central position in the sequence of photosynthetic electron transport carriers, oxidizing plastoquinol (PQH2) and providing the electron transfer connection between the two reaction centre complexes, PSII and PSI, to which H+ transfer is coupled. Therefore, it gives the contribution to transmembrane gradient. Electrons are transferred to PSI via plastocyanin or cyt c6 (Bio.purdue.edu). The proton-pumping cytochrome b6f complex has been shown by biochemical and mass spectroscopic analysis to be a dimer and to contain 8 tightly bound subunits per monomer in the cyanobacterium Mastigocladus laminosus, and 9 in plant chloroplasts (Gómez et al., 2002), (Whitelegge et al., 2002). The mass of the dimer in cyanobacteria is 217 kDa, which indicates that this complex is substantial in size, but somewhat smaller than PSI and PSII. One of the interesting issues is where the cytochrome b6f complex is located in the photosynthetic membrane in vivo and how it cooperates with the photosystems. One possibility includes connection of the cytochrome b6f complex with PS I. Both PSI and PSII can make supercomplexes with LHCII. There is evidence that cytochrome b6f may bind the transition particle of photosystem I – LHCII (Iwai et al., 2010). The study of characterizing such a supercomplex is just in the beginning.

Prokaryotes: classification and photosynthesis Historically, prokaryotes were the first organisms on the Earth which appeared about 3.5 billion years ago. First prokaryotes were adapted to the extreme conditions of early earth. Prokaryotes are divided taxonomically into two domains: the archaebacteria (or archae) and the eubacteria. Despite the fact that some archaebacteria can stand extreme conditions (high temperature, low pH), there are no archaeal species known that carry out chlorophyll-based photosynthesis (Blankenship, 2014). There are only eubacterial species with photosynthetic capacity, although a vast majority of prokaryotes are non-photosynthetic organisms. As of September 2012, there are 30 phyla in the domain "Bacteria" accepted by LPSN, List of Prokaryotic Names with Standing in Nomenclature (Euzéby, 1997), (Parte, 2014). According to the taxonomy division among prokaryotes there are 6 phyla of bacteria that could photosynthesize (Garrity and Holt, 1997),

(Chapman, 1946), (Hohmann-Marriott and Blankenship, 2011). Out of the 6 phyla, 5 are able to carry out anoxygenic photosynthesis. They do not produce oxygen as a by-product of photosynthesis (Woodbury and Allen, 1995). Water is therefore not used as an electron donor. These groups are the purple bacteria, the green sulphur bacteria, the green nonsulphur bacteria, the heliobacteria and the chloroacidobacteria. The remaining sixth phylum can perform oxygenic photosynthesis. This group of bacteria consists of the cyanobacteria (Grossman et al., 1994). A further common division in bacteria is based upon differences in the structure of the cell wall. Considering of the prokaryotic cell bacteria can be divided of gram-positive and gram-negative. In Gram-negative bacteria, including most types of phototrophic bacteria, a second, more permeable, outer membrane is present, as well as a rigid cell wall that provides mechanical stability (Madigan, 2012). This division is, however, not often considered in photosynthetic organisms. For instance, RNA trees place the heliobacteria among the Firmicutes, considered to be gram-positives, but they do not stain gram-positively.

Purple bacteria Purple bacteria or purple photosynthetic bacteria are proteobacteria that are phototrophic. This means they can produce their energy through photosynthesis (Bryant and Frigaard, 2006). The majority contains bacteriochlorophyll a (BChla) and a few others have bacteriochlorophyll b. BChl a and BChl b have wavelengths of maximum absorption in ether solution at 775 nm and 790 nm, respectively. In vivo however, due to shared extended resonance structures, these pigments were found to maximally absorb wavelengths out further into the near-infrared. Other families of photosynthetic bacteria contain other types of bacteriochlorophyll. The green sulphur bacteria contain bacteriochlorophyll c, d, or e, in addition to BChl a and chlorophyll a. Purple bacteria have also various carotenoids, that together in combination with BChl determine their colour – from purple to even green. Based on the range of abilities to metabolize reduced sulphur compounds the purple bacteria can be divided into two groups – sulphur and nonsulphur (Frigaard and Dahl, 2008). However, the terms sulphur and nonsulphur are somewhat misleading because all purple bacteria have the capability to carry out extensive sulphur metabolism (Blankenship, 2014). Most purple bacteria are also capable of nitrogen fixation.

Photosynthesis takes place at reaction centres (RC) that are located in a specially modified portion of the inner cell membrane called the intracytoplasmic membrane, that sometimes is folded into tubes, vesicles, or

A B

Figure 5. Schematic model of the RC-LH1 core complex from Rhodopseudomonas palustris with a central RC surrounded by 15 LH1 units. Transmembrane helices are drawn as ribbons with the program RIBBONS (Carson, 1997). (A) View of the complex perpendicular to the membrane plane. The 5 α- helices of the L subunit (red) and M subunit (purple) of the RC bind a number of bacteriopheophytin and bacteriochlorophyll pigments (green), where charge separation takes place. In addition, the RC has a single α- helix of subunit H. Each LH1 unit is composed of two α-helices of the 훼 and 훽 apoproteins (green and bright blue) with two bacteriochlorophylls (red) in between. The helix W (dark red) keeps the LH1 ring open, to allow diffusion of small electron carriers to the RC. (B) Narrow section of the complex viewed parallel to the membrane plane. Source: (Roszak et al., 2003) with modifications. flat lamellar membranes to increase the available surface area. The purple bacterial reaction centre from Blastochloris viridis (old name Rhodopseudomonas viridis) was crystallized in 1985 (Deisenhofer et al., 1985). It was the very first membrane protein for which a structure was solved. The RC of purple bacteria is build up by three constant membrane protein subunits – L (light), M (medium), H (heavy). Some species have a fourth hydrophilic subunit – C (cytochrome). The L and M proteins in the purple bacterial reaction centre are the functional analogues of D1 and D2 proteins of PSII of the higher plants and also show weak but definite 34

sequence homology to D1 and D2 (Michel and Deisenhofer, 1988) (Fig. 5). The essential photosynthetic steps in purple bacteria occur inside a membrane protein assembly known as the photosynthetic core complex. The core complex consists of a reaction centre surround by a ring of light harvesting complex I (LH1) units. The main function of LH1 is to absorb and transfer the excitation energy of light to the RC. The energy is utilized to create a charge separation difference across the cellular membrane, finally driving proton pumping and conversion of ADP to ATP. LH1 antenna forms a stoichiometric complex with the RC, called the RC- LH1 core complex (Roszak et al., 2003). The structure of the RC-LH1 complex was determined by X-ray crystallography and visualized by atomic force microscopy (Fig. 5) (Roszak et al., 2003), (Sturgis et al., 2009), (Qian et al., 2013). In some species, the RC-LH1 complex is found as a dimer of two closely interacting RC-LH1 complexes, while in other as a monomer. LH1 contains bacteriochlorophyll a (Bchla) and carotenoids that are noncovalently bound to two types of low molecular weight (5 to 7 kD), hydrophobic apoproteins, called α and β, each of which has a single membrane-spanning α helix. The

Figure 6. Different structural organizations of the bacterial photosynthetic core complexes. (A) A core complex in which the LH1 subunits form a complete ring, as present in Rhodospirillum rubrum. (B) A core complex in which the LH1 subunits forms a ring with a gap, with an extra polypeptide near the gap, as in Rhodopseudomonas palustris. (C) and (D) Two proposed organizations for a dimeric core complex, the dimerization of which requires the extra polypeptide, PufX. (C) is drawn according to the highest structural data for a dimeric core complex (Qian et al., 2005), which places PufX near the LH1 gaps. In (D), PufX is assumed to dimerize and is situated at the centre of the core complex. Dimeric core complexes are seen in certain Rhodobacter species, the best-known case being Rhodobacter sphaeroides. LH1 α and β-polypeptides are united for the simplicity and marked in blue color. Picture was adopted from (Ks.uiuc.edu) 35

functioning LH1 complexes are oligomers of these αβ pairs. Each pair or unit binds two bacteriochlorophylls between α and β, together with some other associated pigments (Cogdell et al., 1999). In different bacterial species, the photosynthetic core complex can take on variable types of organizations. Typically, the LH1 subunits surround directly the slightly elliptical RC. In some cases, the LH1 forms a complete ring, an example being Rhodospirillum rubrum (Fig. 6). The core complexes of certain species contain an additional, single transmembrane α-helical protein. In Rhodopseudomonas palustris, this extra protein is called the W protein, and the LH1 ring is seen to exhibit a gap where the W protein resides (Fig. 6, B). In the Rhodobacter species, the LH1 ring around the reaction centre is interrupted by a small protein that breaks the symmetry of the ring and called PufX. This small protein causes the core complex to dimerize in a direct or indirect way, as exemplified by Rhodobacter sphaeroides (Fig. 6, C and D). In some organisms the PufX gene product is lacking and another as yet unidentified protein occupies the same position. The function of these proteins apart from dimerization is thought to provide an opening in the ring that facilitates the diffusion of quinones in and out of the complex (Blankenship, 2014).

Cyanobacteria The cyanobacteria are another phylum of bacteria that obtain their energy through photosynthesis. This are a large and diverse group of photosynthetic prokaryotes (Grossman et al., 1994), (Ho et al., 2011). Cyanobacteria can be found widespread around the world in both terrestrial and aquatic habitats where light is accessible. Most cyanobacteria are photoautotrophs although some species are photoheterotrophs. Most cyanobacteria contain in addition to a plasma membrane specific thylakoid membranes where the photosynthetic apparatus is located (Van De Meene et al., 2006), (Liberton et al., 2011). All cyanobacteria contain chlorophyll a and in the same time most of them fully lack chlorophyll b. Additionally cyanobacteria contain bilin pigments that are organized into large extrinsic macromolecular antenna complexes called phycobilisomes. Phycobilisomes are giant protein complexes with up to 600 polypeptides, anchored to the stromal side of thylakoid membrane. There are many variations to the general phycobilisome structure. Their shape can be hemidiscoidal in cyanobacteria or hemiellipsoidal in red algae.

A schematical description of the hemidiscoidal phycobilisome is presented in Fig. 7. It represents the most common organization in cyanobacteria. The hemidiscoidal phycobilisome complex consists of three types of pigment– proteins known as biliproteins, along with a number of additional proteins known as linkers. The biliproteins are usually of three major types: phycoerythrin, phycocyanin, and allophycocyanin, which differ in protein identity, chromophore type and attachment, and relative location in the architecture of the phycobilisome complex. The biliproteins contain covalently linked bilin chromophores, which are attached via the other linkages to cysteine residues in the proteins. Some of the biliproteins are arranged into six rods, which attach in a fanlike arrangement to a biliprotein core, made of allophycocyanin, that is attached to the stromal side of the thylakoid membrane, usually in close proximity to Photosystem II (Blankenship, 2014). The scheme of figure 7 is a very simple presentation of the actual situation, because each unit of allophycocyanin, phycocyanin and phycoerythrin is composed of a sandwich of two disks each composed

Figure 7. Schematic model of a tricylindrical hemidiscoidal phycobilisome. Source: (MacColl, 1998) with modifications.

of three subunits. It nevertheless shows the way how these hexameric units form the phycobilisome. At the same time the other types of phycobilisomes were found that are transferring the energy to the Photosystem I (Liu et al., 2013), (Watanabe et al., 2014). These novel types of phycobilisomes consist of just single rods in which units of phycocyanin and phycoerythrin are stacked, but without allophycocyanin. Each phycobiliprotein has a specific absorption and fluorescence emission maximum in the visible range of light. The phycobilisome represents a

classic example of the funnel concept in photosynthetic antennas, in that the phycoerythrin at the distal ends of the rods absorbs at the shortest wavelength, while the phycocyanin is intermediate, and the allophycocyanin absorbs at the longest wavelength, or lowest energy. Energy transfer processes within the phycobilisome serve to direct the energy toward the membrane, where it is trapped by photochemistry (Blankenship, 2014). A second level of control is provided by the linker polypeptides, which tune the absorbance maxima of the biliproteins to facilitate the transfer process. Thus, the cells take advantage of the available wavelengths of green light in the 500-650 nm range, which are inaccessible to chlorophyll, and utilize this energy for photosynthesis. This also makes cyanobacteria and plants complementary in fully utilizing light in aquatic habitats. The composition of the phycobilisome can be modified due to the conditions changing, in particular the light intensity. Thus a species lacking phycoerythrin has at least two additional disks of phycocyanin per rod, which is sufficient for maximum photosynthesis (Lea-Smith et al., 2014). This phenomenon is known as chromatic acclimation (Grossman et al., 1993), (Gutu and Kehoe, 2012). A final remark is about the geometrical arrangement of the phycobilisome, which is very elegant and results in 95% efficiency of energy transfer (Glazer, 1985).

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