University of Groningen
Structures of photosynthetic membrane complexes
Semchonok, Dmytro
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Chapter 1.
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 H 2,
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
7
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
discovered by Melvin Calvin's group, who used 14CO2 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
8
one with the lowest catalytic turnover rate (1–3 s–1). 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 lightharvesting 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
9
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.
10
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 = 𝛮𝛢 𝑐𝑜𝑛𝑑𝑒𝑛𝑠𝑒𝑟+𝑁𝐴 𝑜𝑏𝑗𝑒𝑐𝑡𝑖𝑣𝑒;
where R = resolution, measured as distance, depends on the angular aperture
α; NA = numerical aperture, equal to ηsin(𝛼/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.
11
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 = 1.45+ 0.95 = 203 nm;
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:
χ(ν) = 2π/λ (-Cs(λ4ν4/4) + Δf(λ2ν2/2) )
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)):
Sin(χ(ν)) = Sin[2π/λ (-Cs(λ4ν4/4) + Δf(λ2ν2/2) )],
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
14
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
good signal-to-noise ratio for small
molecules
simple to apply
resistant to radiation
works well on heterogeneous preps
can induce preferred orientation
3-D reconstruction is possible
Disadvantages
molecules are prone to structural
collapse
high background from surrounding
stain
possible chemical interactions
distortions due to ionic strength and
low pH
limited resolution
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.
15
Table 2. Advantages and disadvantages of cryo-technique
Advantages
no artefacts due to fixation,
dehydration or staining
preservation of native
conformation
random orientation
good contrast at high defocus
higher resolution info than
negative stain
Disadvantages
low signal-to-noise-ratio
sensitive to radiation
technically challenging
difficult to distinguish between different
orientations vs. conformations
freezing artefacts
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 singleparticle 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.
16
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 rhodopsinbased 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
17
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.
18
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
19
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, PsbPlike, 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.
20
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 lightharvesting 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
21
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).
22
Table 3. Subunit composition of plant and cyanobacterial
Photosystem II
Subunit
name
PSII-A (D1)
Gene
PSII-B
(CP47)
PSII-C
(CP43)
PSII-D (D2)
psbB
C
56
psbC
C
51
psbD
C
39
psbE
C
psbA
Gene
Mass
a
location (kDa)b
C
39
Cofactors
Function
Core reaction
centre of
Photosystem II
9
chlorophyll,
pheophytin,
quinone,
𝛽-carotene, Fe
chlorophyll,
𝛽-carotene
chlorophyll,𝛽carotene
chlorophyll,
pheophytin,
quinone,
𝛽-carotene, Fe
heme
PSII-E
(cytb559𝛼)
PSII-F
(cytb559𝛽)
PSII-H
psbF
C
4
heme
psbH
C
8
phosphate
PSII-I
psbI
C
4
PSII-J
psbJ
C
4
PSII-K
psbK
C
4
PSII-L
PSII-M
PSII-N
PSII-O
(OE33)
psbL
psbM
psbN
psbO
C
C
C
N
4
4
5
27
Core antenna
Core antenna
Core reaction
centre of
Photosystem II
Core reaction
centre of
Photosystem II
Core reaction
centre of
Photosystem II
Photoprotection,
QA to QB
regulation
Core reaction
centre of
Photosystem II
Assembly of
Photosystem II
Role in PSII
assembly
Role in QA binding
Role in PSII stability
Role in PSII stability
Stabilizes Mncluster, Ca2+
and Cl− binding
23
PSII-P
(OE23)c
PSII-Q
(OE16)c
PSII-Rc
PSII-S
(CP22)
PSII-T
(ycf8)
PSII-Ud
PSII-V
(cytc-550)d
PSII-Wc
PSII-X
PSII-Y
(ycf32)
PSII-Z
(ycf9)
PsbP
N
20
Ca2+ and Cl− binding
PsbQ
N
17
Ca2+ and Cl− binding
PsbR
PsbS
N
N
10
22
psbT
C
3
?
Antenna regulation
by
xanthophyll cycle
Role in PSII stability
psbU
psbV
14
15
chlorophyll,
carotenoids
heme
Role in O2 evolution
Role in O2 evolution
PsbW
psbX
PsbY
N
C
N
6
4
4
Role in PSII stability
Role in QA function
Mn binding?
psbZ
C
9
Psb27
slr1645
12
Psb28
sll1398
13
Psb29
sll1414
27
Psb30
(ycf12)
Psb31e
Psb32
LHCIIouterc
LHCIIouterc
LHCIIaouterc
LHCIIinner
(CP29)c
sll0047
3
Antenna-reaction
centre
interaction
Role in PSII
assembly
Role in PSII
assembly
Role in PSII
assembly
Stabilize PSII
sll1390
Lhcb1
N
12
22
30
Role in O2 evolution
Role in PSII repair
Antenna function
Lhcb2
N
31
Lhcb3
N
25
Lhcb4
N
35
chlorophyll,
carotenoids
chlorophyll,
carotenoids
chlorophyll,
carotenoids
chlorophyll,
carotenoids
Antenna function
Antenna function
Antenna function
24
LHCIIinner
(CP26)c
Lhcb5
N
36
chlorophyll,
carotenoids
Antenna function
LHCIILhcb6
N
18
chlorophyll,
Antenna function
inner
carotenoids
(CP24)c
a
Gene location applies only to eukaryotic organisms. C, chloroplast; N, nucleus
b
Mass is actual mass based on gene sequence
c
Found only in eukaryotic organisms
d
Found only in cyanobacteria
e
Found 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
25
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).
26
A
B
C
D
E
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 from
27
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
name
Gene
name
Gene
locationa
Mass
(kDa)b
PSI-A
(PsaA)
psaA
C
84
PSI-B
(PsaB)
psaB
C
83
PSI-C
psaC
C
9
PSI-D
PsaD
N
18
PSI-E
PsaE
N
10
PSI-F
PsaF
N
17
PSI-Gc
PSI-Hc
PsaG
PsaH
N
N
11
11
PSI-I
PSI-J
PSI-K
psaI
psaJ
PsaK
C
C
N
4
5
9
PSI-L
PsaL
N
18
PSI-Md
psaM
PSI-Nc
PsaN
PSI-Xd
psaX
LHC-Ic
(LHCI720)
LHC-Ic
Lhca1
Lhca2
Cofactors
Function
chlorophyll,
quinone,
𝛽-carotene,
Fe–S
chlorophyll,
quinone,
𝛽-carotene,
Fe–S
Fe–S
Core reaction
centre of PSI,
charge separation,
electron transport
Core reaction
centre of PSI,
charge separation,
electron transport
Role in electron
transport
Ferredoxin
docking
Role in cyclic
electron transport
Plastocyanin
docking
Role in QA binding
Interaction with
LHCII
?
?
Role in docking of
LHCI
Trimer formation
in cyanobacteria
Trimer formation
in cyanobacteria
Plastocyanin
docking
Binding of IsiA
antenna
Antenna function
3
N
10
3
chlorophyll
N
22
chlorophyll,
carotenoid
N
23
chlorophyll,
Antenna function
28
(LHCIcarotenoid
680)
LHC-Ic
Lhca3
N
25
chlorophyll, Antenna function
(LHCIcarotenoid
680)
LHC-Ic
Lhca4
N
21
chlorophyll, Antenna function
(LHCIcarotenoid
720)
a
Gene location applies only to eukaryotic organisms. C, chloroplast; N,
nucleus
b
Mass is actual mass based on gene sequence
c
Found only in eukaryotic organisms
d
Found 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.
29
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 membraneassociated 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 ctype 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
30
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 (lightblue 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).
31
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),
32
(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.
33
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 αhelices 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 αhelix of subunit H. Each LH1 unit is composed of two α-helices of the 𝛼
and 𝛽 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 RCLH1 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.
36
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
37
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).
38
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