light-dependent reactions of photosynthesis...when only one reaction center is present,...
TRANSCRIPT
TEACHING TOOLS IN PLANT BIOLOGY™: LECTURE NOTES
Light-Dependent Reactions of Photosynthesis
INTRODUCTION AND OVERVIEW
Photosynthesis in plants converts the energy of sunlight into
chemical energy and through this process releases O2. Although
photosynthesis involvesmanyproteinsandcatalyticprocesses, it
often is described as two sets of reactions, the light-dependent
reactions and the carbon-fixing reactions, which are also occa-
sionally described as the “dark reactions.”
Plants do not have a monopoly on photosynthesis, which also
occurs in other photosynthetic eukaryotes and several species
of bacteria. In most cases, bacterial photosynthesis does not
release oxygen. Oxygenic (O2-producing) photosynthesis is
restricted to cyanobacteria, green, red, and brown algae, and
plants. Oxygen is produced when water is split to release
protons (H1), electrons, and oxygen. Oxygenic photosynthesis
requires the involvement two photosystems working in series,
photosystem I (PSI) and photosystem II (PSII).
The light-dependent reactions of photosynthesis in plants can
bedescribed as a series of steps that take place in and across the
thylakoid membranes of chloroplasts. Light harvesting involves
the capture of photonsbypigmentmolecules and the funneling of
light energy to reaction center chlorophyll molecules. Reaction
center chlorophylls are positioned so that the captured light
energy induces a charge separation across the membrane. Light
excitation of the reaction center chlorophyll of PSII causes it to
give up electrons and become oxidized. Electrons stripped from
water with the accompanying production of oxygen reduce the
oxidized reaction center chlorophyll in PSII. Electrons are ulti-
mately passed by way of the electron transport chain of cyto-
chrome b6f and through PSI to NADP1, reducing it to NADPH.
During these reactions, protons are translocated to the interior
lumen compartment of the thylakoid membranes. The resulting
charge and pH gradients, collectively known as proton motive
force, drive the synthesis of ATP by the ATP synthase complex.
This lesson examines how the multiprotein complexes that
make up the photosynthetic apparatus carry out these steps,
their evolutionary origins, and how their inherent flexibility and
feedback loops enable photosynthetic reactions to acclimate to
changing light andmetabolic conditions. Finally, we explore how
current insights are enabling scientists to develop improved algal
strains for biofuels and plants that are more resilient to abiotic
stresses and are inspiring the design of artificial systems for the
conversion of sunlight into energy.
Our understanding of photosynthesis rests on the work of
thousands of scientists working in physics, engineering, and
chemistry and across the disciplines of biology. Pioneers include
JosephPriestley, who recognized the ability of plants to “restore”
air that had been “injured” by a burning candle (in other words,
to produce oxygen). Key discoveries came from the work of
Willstatter (Nobel Prize in Chemistry, 1915), Warburg (Nobel
Prize in Physiology or Medicine, 1931), Hill (1930s), Emerson
(1950s), Arnon (1950s–1970s), Calvin (Nobel Prize in Chemistry,
1961), Woodward (Nobel Prize in Chemistry in 1965 for the total
synthesis of chlorophyll and other molecules), Deisenhofer,
Huber, and Michel (Nobel Prize in Chemistry in 1988 for the
determination of the three-dimensional structure of a bacterial
photosynthetic reaction center), and others too numerous to
mention. For comprehensive histories of the pioneers of pho-
tosynthetic research, seeGovindjee andGest (2002), Govindjee
et al. (2003), and Govindjee et al. (2004).
EVOLUTION AND DIVERSITY OF PHOTOSYNTHESIS
It is difficult to set a date for the beginning of photosynthesis.
We knowwith certainty that there is a layer of insoluble iron that
was deposited in rocks ;2.3 to 2.5 billion years ago, which
resulted from a dramatic increase in the level of atmospheric
oxygenand theconcomitantdecrease in solubility of irondue to
its oxidationbyoxygen to the formof iron(III) oxide. This sets the
most recent date for the evolution of oxygenic photosynthesis,
but evidence from stromatolites (ancient fossils) suggest that
oxygen-producingcyanobacteriamayhave evolved longbefore
this, perhaps as long as 3.4 billion years ago. The time lag
between the potential origin of oxygenic photosynthesis and
the accumulation of atmospheric oxygen that occurred millions
of years later can be attributed largely to the ability of oceans to
accumulate and buffer oxygen. Suffice it to say, photosynthesis
has beenmaking a tremendous contribution to the atmospheric
gas composition and the entry of energy into Earth’s biosphere
for a long time.
Although photosynthetic prokaryotes are found in several dif-
ferentphyla, there arebasically threedifferent typesofchlorophyll-
based photosynthesis. Anoxygenic photosynthetic prokaryotes
use either a Type I or a Type II reaction center (RC), described
further below. As the name indicates, these organisms do not
oxidizewaterbut insteaduseanelectrondonorother thanwater.
Oxygenic photosynthesis, which is restricted to cyanobacteria
and the eukaryotic organisms that have cyanobacteria-like
chloroplasts, requires both Type I and Type II reaction centers
working in series to bridge the energy gap between water
oxidation and NADP1 reduction.
The evolution of photosynthesis has been described as,
“a complex process that cannot be described by a simple, linear,www.plantcell.org/cgi/doi/10.1105/tpc.15.tt1115
The Plant Cell, November 2015, www.plantcell.org ã 2015 American Society of Plant Biologists. All rights reserved.
branching evolutionary diagram.Rather, photosynthesis emerged
by recruiting and modifying genes encoding components of
several other pre-existing metabolic pathways, along with a
few key innovations and probably several lateral gene-transfer
events. The resulting view is that, likemanymetabolic pathways,
photosynthesis is a mosaic process that has no single well
defined evolutionary origin,” (Blankenship, 2001).
Anoxygenic Photosynthesis
Photosynthetic bacteria have been found in six phyla, of which
five contain only anoxygenic photosynthetic bacteria. The wide-
spread occurrence of photosynthesis and the fact that only some
members of these phyla are photosynthetic lends support to the
idea that lateral gene transfer (the direct movement of genes
across species, not via a common ancestor) has been a major
factor in the distribution of photosynthetic capacity.
Anoxygenic photosynthetic prokaryotes use either a Type I or
Type II reaction center (and it is likely that Type II reaction centers
are derived from Type I). Both types share the fundamental
property of using light energy to induce charge separation
across a membrane using bacteriochlorophyll as the primary
electron donor. They differ in the first stable electron acceptor,
which in Type I RCs are Fe4-S4 clusters and in type II RCs are
quinone molecules.
When only one reaction center is present, photosynthetic
electron transfer is cyclic and the light-driven reaction mainly
promotes the establishment of a proton motive force for the
production of ATP.
Oxygenic Photosynthesis
Cyanobacteria
Cyanobacteria are a monophyletic group but are also among
the most morphologically diverse and successful prokaryotes.
They are important contributors to Earth’s oxygen supply and
the conversion of solar energy to chemical energy. Many cya-
nobacteria are also able to fix nitrogen (convert chemically inert
N2 into NH41, a form that can be used by organisms). Most
cyanobacteria are free-living organisms in aquatic ecosystems.
Some cyanobacteria livewithin plant tissues as endophytes and
can contribute to the plant’s nitrogen nutrition, and some form
symbiotic associations with fungi in the form of lichen or even
with marine animals (see below). Cyanobacteria provide an
outstanding model for the study of oxygenic photosynthesis.
Eukaryotic Photosynthesis: Primary and Secondary
Endosymbiosis
With a single known exception (Paulinella chromatophora;
describedbelow), photosynthetic eukaryotes aredescendants
of a primary endosymbiotic event that took place ;1 to 1.5
billion years ago, in which an ancestral eukaryotic cell engulfed
anancestral cyanobacterium.Thisprimaryendosymbioticevent
resulted in three lineages: Viridiplantae (green algae including
Chlamydomonas reinhardtii and land plants), rhodophytes (red
algae), and glaucophytes (rare freshwater algae).
In themillions of years since this primary endosymbiotic event,
many of the endosymbiont’s genes were transferred to the
nucleus, making photosynthetic function dependent on both
nuclear and organelle genomes. Many plastid proteins are
encoded by nuclear genes; their genes are transcribed in the
nucleus, their mRNAs are translated in the cytosol, and the
resulting proteins are transported into the plastid through
a two-layered envelope that resembles the cyanobacterial
inner and outer membranes. The development of machinery
for the import of proteins from cytosol to plastid was essential
for the transition from free-living cyanobacteria to plastid. The
selective advantage for genemigration from endosymbiont to
nucleusmaybe that thenuclear locationof thesegenesprovides
a better integration of their function or because nuclear genes,
unlike plastid-localized genes, can self-correct during meiosis
via recombination.
Although green algae and terrestrial plants descend from the
same ancestor, there have been some changes in their photo-
synthetic machinery since they diverged;400million years ago.
The core reaction center complexes of PSI and PSII are more or
less conserved among cyanobacteria, algae, and plants, but the
light-harvestingcomplexes (LHCs) that capturephotons toenergize
PSI and PSII differ in different photosynthetic organisms.
Finally, secondary and tertiary endosymbiosis has extended
the ability to carry out photosynthesis much further than the
progeny of the primary endosymbiotic event. Secondary and
tertiary endosymbiosis refers to the process in which green or
red algae (the products of primary endosymbiosis) were them-
selves engulfed within other cells. Products of these events
include euglenoids, diatoms, dinoflagellates, and other algae.
Collectively, these organisms belong to a broad spectrum of
eukaryotic clades, and although they are important in global
energy transfer, they are less commonly used asmodels for the
study of light-dependent photosynthetic reactions.
Nascent Primary Endosymbiosis in P. chromatophora
Recently, a second example of primary endosymbiosis was
identified in thephotosynthetic amoebaP.chromatophora,which
is in the kingdom Rhizaria. This second primary endosymbiotic
event occurred relatively recently, ;60 million years ago; recall
that the primary endosymbiotic event that gave rise to plants
occurredmore thanabillionyearsago.P.chromatophoraharbors
photosynthetic organelles called chromatophores, which resem-
ble plastids. Chromatophores in P. chromatophora are derived
from a-cyanobacteria, whereas all other known plastids in pho-
tosynthetic eukaryotes are derived from b-cyanobacteria. The
chromatophore is clearly an endosymbiotic organelle because it
cannot live independently outside of the host and it divides with
the host cell. Like plant plastids, some of the P. chromatophora
genes have been transferred to the nucleus, although to a lesser
extent as those in chloroplasts probably due to their younger
evolutionary age. As in plants and green algae, the proteins
encoded by these transferred nuclear genesmust be transported
into the organelle. This exciting discovery offers researchers
a system in which to explore an early stage of plastid evolution
and ultimately to understand better the evolution of eukaryotic
photosynthetic organelles.
2 The Plant Cell
LIGHT AND PIGMENTS
The Nature of Light
Photosynthesis converts light energy into useful biochemical
products. Light is composed of energy-carrying particles known
asphotons. Photons can alsobe represented as electromagnetic
waves that travel at a fixed speed of;3 3 108m/s (denoted as c)
with a frequency (n; cycles per second) andwavelength (l; dis-
tance between peaks) in the relationship c 5 n l. Since c is
a constant, as l decreases, n increases; the shorter the wave-
length, the higher the frequency. Furthermore, since E 5 hn
(whereE is a photon’s energy, h is Planck’s constant, and n the
photon’s frequency), as wavelength decreases, frequency
increases and so does energy. Long-wavelength light such
as that in the infrared region of the spectrum is low in energy,
whereas short-wavelength light such as that in the UV region
of the spectrum carries much more energy. Thus, the wave-
length of light determines whether or not it has enough energy
to drive the reactions of photosynthesis. In plants, PSII has an
energy requirement for light of 680 nm or shorter, and PSI
requires light of 700 nm or shorter.
Visible light is defined by the properties of the human eye and
falls between ;400 and 700 nm. Electromagnetic radiation of
wavelengths less than 400 nm include high-energyUV light and
x-rays, which are largely blocked from reaching the earth’s
surface by its atmosphere. Electromagnetic radiation of wave-
length longer than 700 includes low-energy infrared light and
radio waves.
Photosynthetic Pigments and Accessory Pigments
Pigments are compounds that absorb visible light and are often
characterized by networks of double bonds. As a consequence
of the pigments they produce, plant chloroplasts preferentially
absorb blue and red light, which is why leaves and other pho-
tosynthetic tissues of plants appear green. When a pigment
molecule’s conjugated electron system absorbs light, an elec-
tron becomes excited to a higher energy state that can be
passed on to anothermolecule. In photosynthetic systems, light
is absorbed by accessory pigments present in protein-pigment
arrays arranged in LHCs. A system of accessory pigments sur-
rounds each reaction center in what is described as an antenna
complex. Each antenna complex passes the energy of collected
light to a reaction center at its center. The size and composition of
the antenna complex is sensitive to the environment; for example,
a heavily shaded leaf has a larger antenna complex surrounding
each reactioncenter thana leaf in full sunlight.Accessorypigments
include chlorophylls, carotenoids, and phycobilins (in cyanobac-
teria and red algae).
Chlorophyll
Most chlorophyll-dependent photosynthetic organisms produce
chlorophyll a, which is always present in the reaction center,
and one or more additional forms of chlorophyll that serve as
accessory pigments. Chlorophyll comprises a cyclic tetrapyr-
role moiety with a central Mg ion and an acyl (lipid) tail through
which it is anchored. Chlorophyll is produced in plastids through
the tetrapyrrole biosynthetic pathway, which is a branched
pathway that also leads to heme, siroheme, and phytochromo-
bilin, which is a linear tetrapyrrole that is the chromophore of the
photoreceptor phytochrome. Structurally related to chlorophyll
are pheophytins that are the first electron acceptor from the
excited chlorophyll of PSII. Pheophytin is simply chlorophyll
without the Mg.
Substitutions on the tetrapyrrole ring structure affect chloro-
phyll’s absorption spectra. Chlorophyll b is found in land plants,
green algae, and cyanobacteria. Chlorophyll c lacks the phytol
tail and is primarily found in brown algae and dinoflagellates.
Chlorophyll d, e, and f are minor components and found in
diverse organisms, mostly cyanobacteria. Bacteriochlorophylls
are structurally similar to plant chlorophylls and are produced by
a conserved biosynthetic pathway. Plant chloroplasts produce
chlorophyllaandb,which show twopeaksof light absorbance in
blue and red light. Chlorophyll a absorbsmaximally at;430 and
660 nm, and chlorophyll b at ;460 and 650 nm, with peak
absorbance wavelength varying slightly with solvent. Chloro-
phyll a and b are differentially represented in core reaction
centers and antennacomplexes,so therelativeabundanceofeach
is an indicator of the proportional representation of each type of
complex.
When chlorophyll a at the reaction center absorbs light energy,
an electron moves to a higher energy level, and the pigment
is denoted as Chl*. The excited electron can fall back to its
lower energy level, releasing energy as longer wavelength
light (fluorescence), the energy can be released as heat, or the
electron can be transferred to another molecule as the first
step of photochemistry; when this happens, the resulting
chlorophyllmolecule is denoted asChl1. The rest of this article
describes what happens after the charge transfer event that
initiates photochemistry.
Carotenoids
Carotenoids are accessory pigments that absorbmainly in the
blue range and appear yellow or orange; they capture photons
that would not be absorbed by chlorophyll and pass the
energy to the reaction center chlorophyll. Carotenoids have
essential functions in photoprotection, including the dissipa-
tion of excess energy when light levels exceed the plant’s
ability to process it, and they also structurally stabilize the
photosystems.
There are two classes of carotenoids: hydrocarbon carotenes
andoxygen-containingxanthophylls.Bothare40-carbon lipophilic
pigments made from isoprenes. Carotenes such as a-carotene,
b-carotene, and lycopene appear orange; they are named after
the orange color of carrots but are also found in sweet potato,
pumpkin, and other orange-colored tissues. Xanthophylls
appear yellowish (xanthos in Greekmeans yellow) and include
lutein, zeaxanthin, and violaxanthin; the latter will be discussed
later in the context of the xanthophyll cycle, which is involved in
nonphotochemical quenching (NPQ) of excess light energy. Ca-
rotenoids are important nutrients for humansaswell;b-carotene is
the precursor for vitamin A, and lutein accumulates in the retina
November 2015 3
of the eye where it protects photopigments from damage. Carot-
enoids are abundant in photosynthetic tissues but their color is
masked by that of chlorophyll; when chlorophyll breaks down
during senescence, they are revealed as the familiar yellow and
orange colors of autumn leaves.
Phycobilins
Phycoblilins are linear tetrapyrroles that comprise the chromo-
phore moiety of phycobiliproteins, which are water-soluble
accessory photosynthetic proteins in cyanobacteria, red algae,
and other non-green algae. In cyanobacteria, phycobilins are
found in phycobilisomes, which act as light-harvesting antenna.
Thephycobilins areespecially efficient at absorbing red, orange,
yellow, and green light, wavelengths that are not well absorbed
by chlorophyll a. Small modifications to the phycobilin chromo-
phore or to their orientation relative to the apoprotein affect phy-
cobiliprotein absorption spectra. Phycobilin pigments include
phycocyanin (abundant in cyanobacteria, absorbs red and orange
light, and appears blue) and phycoerythrin (abundant in red algae,
absorbs blue and green light, and appears red). The chromophore
of phytochrome, which regulates light responses, is a linear
tetrapyrrole related to phycobilins.
THE LIGHT RESPONSE CURVE AND QUANTUM
EFFICIENCY
The light response curve is an important tool for describing the
rate of photosynthesis and is produced by plotting the rate of
photosynthesis (measured as O2 evolution or CO2 consumption)
at varying light intensities. At low light levels, photosynthesis is
limited by light, so there is a linear increase in photosynthesiswith
increasing light. However, this linear relationship only holds at
fairly low light levels. The light saturation point is the light level at
which the rate of photosynthesis is limited by factors other than
light; it depends on many factors including the species of plant,
the light intensity towhich the plant has acclimated, temperature,
etc. In thedarkor at very low light levels,mitochondrial respiration
can be observed as a net consumption of O2 (or production of
CO2). The light level at which respiration and photosynthesis
balance and net O2 production is zero is defined as the light
compensation point; it also depends onmany factors including
the species of plant, temperature, etc.
Quantum efficiency measures the energetic efficiency
of photosynthesis. The quantum yield of oxygen evolution
describes the amount of oxygen evolved per photon. Theo-
retically, it requires eight photons to evolve onemolecule ofO2
(as described further below), so the theoretical quantum
efficiency is 0.125 mol O2/mol photons. The measured value
is always lower, as there are inevitably energy losses due to
heat production, fluorescence, and other factors. Quantum
efficiency is a useful indicator of stress, for example, as stress
induces nonphotochemical quenching, which lowers quan-
tum efficiency. The quantum efficiency is not very dependent
on light wavelength as long as the light has sufficient energy to
drive photosynthesis. Although plants do not absorb much
green light, it is capable of initiating photochemistry.
PLASTIDS AND CHLOROPLASTS
Plastids are essential organelles in most plant cells although
generally not found in the spermcells of pollen. Severalmetabolic
processes besides photosynthesis occur in plastids including
nitrogen and sulfur assimilation and the synthesis of secondary
metabolites, pigments, fatty acids, and various hormones. It is
important to remember that plastids are descendants of endo-
cytosed prokaryotes. Many features of the ancestral prokaryote
can still be detected, including the vestigial circular plastid
genome, division by fission, and the chemical composition of
the lipid membranes.
Chloroplast Development and Differentiation
Plastids are never synthesized de novo but are always inherited
by daughter cells during cell division. Some single-celled algae
have only a single chloroplast. Plant cells have more; undifferen-
tiated meristematic cells may have as few as 10 proplastids that
are small and have minimal expansion of internal membranes,
whereas leaf mesophyll cells may have 100 or so chloroplasts.
Proplastids differentiate into several forms depending on the
cell type. In some root cells, proplastids differentiate into amy-
loplasts thataccumulatestarchgranulesandhaveaspecial role in
gravity perception. In fruits and flowers, some plastids differen-
tiate as carotenoid-filled chromoplasts. In the prephotosynthetic
tissues of dark-grown or etiolated seedlings, plastids partially
differentiate into etioplasts, which are primed to differentiate into
chloroplasts upon illumination. Protochlorophyllide, the precur-
sorof chlorophyll, is attached to theprolamellar body,which is the
thylakoid precursor in etioplasts.
When etioplasts are exposed to light, within just a few hours
there is a light-induced conversion of protochlorophyllide to
chlorophyllide and then to chlorophyll, and theprolamellar bodies
reorganize into thylakoids. Light also triggers changes in gene
expression in the nucleus. Nuclear genes encoding photosyn-
theticapparatusare induced, asare thegenesencodingenzymes
required forpigmentsynthesis.These transcriptional changesare
accompanied by large-scale metabolic changes to prepare the
organelle for its role as a photosynthetic entity.
Ultrastructure of Chloroplasts
Chloroplasts of higher plants are typically oval in shape and;5 to
10 mm long. They are bounded by an outer layer that is a double
membrane (meaning that it has two lipid bilayers) known as
the envelope. Inside the envelope is an aqueous layer called the
stroma, which is largely filled by the photosynthetic membranes
called thylakoids. These membranes anchor the photosynthetic
apparatus and are needed for the establishment of a proton
gradient to drive ATP synthesis. The space inside the thylakoid
membranes is known as the thylakoid lumen. Plastoglobules are
lipoprotein particles permanently attached to the thylakoid mem-
branes, which contain biosynthetic enzymes responsible for the
production of tocopherols (reactive oxygen scavengers, also
known as vitamin E) and phylloquinone (polycyclic aromatic
ketones, part of the electron transport chain of PSI and also
4 The Plant Cell
known as vitamin K1). Of the 3500 to 4000 different proteins
found in the chloroplast, only ;75 to 80 are encoded by the
plastid genome; genes encoding all the rest are located in
the nucleus.
Chloroplast Envelope
Chloroplasts are enclosed by a double membrane called an
envelope. Lipid composition, particularly enrichment in galac-
toglycerolipids, and the presence of distinctly prokaryote-like
membrane proteins indicate that both the outer and inner
envelope membranes are derived from the outer and inner
membranes of ancestral prokaryote. Furthermore, the lipid
composition of the inner envelope membrane is similar to that
of the thylakoid membrane. Membrane proteins in the envelope
include protein import machinery through which the photosyn-
thetic complex proteins are translocated, as well as numerous
transporters for the metabolites that move into and out of a
functioning chloroplast.
Thylakoid Membranes
The inner structures of land plant chloroplasts are beautiful
and distinctive. Thylakoids serve as anchors for large protein
complexes of light-dependent reactions and serve as barriers
to generate the proton gradient required for ATP synthesis.
Undifferentiated plastids have little or no thylakoid mem-
branes. Upon light stimulation, internal thylakoid membranes
proliferate, forming discs of membranes known as grana that
resemble stacks of coins and are connected by regions of
membrane known as the stromal thylakoid. Some regions of
the thylakoid membranes are pressed against each other and
known as appressed membranes. PSII is mainly found in the
appressed regions of themembrane. Themargins of the grana
and the stromal membranes are not pressed against other
membranes and are therefore known as unappressed mem-
branes. The bulkier protein complexes PSI and ATPase predom-
inate on the unappressed membranes, and cytochrome b6f is
uniformly distributed throughout. This physical separation
of photosystems allows a greater fine-tuning of photosynthe-
sis and responses to changing light intensity and metabolic
demands.
STRUCTURE AND FUNCTION OF PHOTOSYNTHETIC
COMPLEXES
Each of the photosynthetic apparatus described below aremem-
brane anchored and comprise multiple proteins and pigments or
cofactors. Classically, the components of these complexes were
characterized by their physical separation through density-
gradient centrifugation and gel electrophoresis, and these
methods are still important. In recent years, our understanding
of photosynthesis has been greatly enhanced through high-
resolution structural analysis based on crystallography of the
key components. Comparisons of the structures obtained
from plant, algae, and cyanobacteria reveal insights into
how evolution has shaped these complex machines.
Linear Electron Transport
Here, we describe the machinery that carries out the light
reactions of photosynthesis in plants, initially focusing on the
linear electron transport (LET) that predominates in most con-
ditions. In LET, light energy transferred from the LHCs to PSII
and PSI oxidizes their reaction centers, driving the linear transport
of electrons fromH2O to NADPH. Specifically, electrons flow from
water through PSII to the mobile electron carrier plastoquinone
(PQ), through cytochrome b6f, then to PSI by another mobile
electron carrier plastocyanin (PC), then via ferredoxin to reduce
NADP1 to NADPH.
Concomitant to the reduction of NADP1 to NADPH, the elec-
tron transfer events also contribute to a proton gradient across
the thylakoid membrane, which is used for the synthesis of ATP
by ATP synthase. Together, ATP and the reducing power of
NADPH are used in the biosynthetic reactions of the carbon-
fixing reactions of photosynthesis.
Structure and Function of PSII-LHCII Supercomplex
PSII carries out light-energized electron transport that leads
to water splitting and oxygen release. PSII is a multi-protein,
multi-pigment complex that spans the thylakoid membrane.
PSII operationally includes the core PSII complex along with
the oxygen-evolving complex (OEC) that is highly conserved
between plants, algae and cyanobacteria, and the peripheral
antenna or light-harvesting complexes (LHCII) that are more
divergent.
Core PSII Complex
The structure of PSII from cyanobacteria has been resolved at
1.9A,whichallowsmostoftheproteinsandcofactorstobeidentified.
Thecyanobacterialcomplex ismadeupof17membrane-spanning
proteins, three peripheral proteins, 35 chlorophyll molecules,
and 11 b-carotenes. Genes encoding PSII proteins are named
Psb genes.
PSII functions as a dimer. Themost highly conserved elements
are those in the reaction center involved in light capture and
electron transport as well as those involved in water splitting.
Eachmonomer has a distinct reaction centermade upof proteins
D1 (PsbA) and D2 (PsbD) as well as an inner light-harvesting
complex composed of chlorophyll binding proteins CP43 (PsbC)
and CP47 (PsbB). The function of the reaction center proteins is
mainly to anchor the pigments and electron carriers in place and
to provide an appropriate environment for efficient energy trans-
fer. As described below, the D1 protein of PSII is subject to light-
dependent turnover and repair, a property that may provide
a safety valve for the protection of other sensitive photosynthetic
components.
A pair of chlorophyll molecules forms the core of the reaction
center. Light excites chlorophyll to form Chl*. Charge transfer
occurswhenanelectron leavesChl*, formingChl1. Theelectron is
passed to pheophytin (chlorophyll that lacks Mg; Pheo) to pro-
duce Pheo2. Pheophytin passes the electron to the primary
acceptor, QA, which is a tightly bound PQ. PSII contains two
(possibly three) PQ cofactors. One, QA, is fixed within the PSII
November 2015 5
structure andservesasaconduit for electrons frompheophytin to
the second PQmolecule. The second PQmolecule binds to PSII
at theQB site. Plastoquinone is a two-electron carrier, which after
accepting twoelectrons is in the formPQ22. Twoprotons fromthe
stroma bind to it to form PQH2 (plastoquinol), an uncharged
molecule. This reduced, uncharged form can dissociate from
thePSII complex, physically carryingelectrons to thecytochrome
b6f complex and simultaneously transferring protons from the
stroma to the lumen.
A pool of PQ/PQH2 molecules exists within the thylakoid
membrane to shuttle electrons between PSII and cytochrome
b6f. Once PQH2 is released from PSII, another PQ molecule will
bind to PSII in its place to accept electrons from another round of
excitation. It should be noted that PSII can only give rise to one
charge separation event at a time. Thus, twophotonsare required
for the complete reduction and release of a single PQH2molecule
from the complex.
Extrinsic Proteins and the Oxygen-Evolving Complex
The OEC resides on the luminal surface of PSII and includes the
catalytic center, loops of intrinsic proteins, and some extrinsic
proteins. There is one OEC per PSII monomer. The catalytic
center core is an inorganic Mn4CaO5 cluster that performs the
mechanistically challenging reaction of removing four tightly
bound electrons and four protons from water to form O2. The
cluster cancycle throughdifferent oxidationstates tosequentially
provide four electrons and reach the oxidation status required to
split two water molecules and release O2. The four H1 released
from the water molecules on the luminal side of the membrane
contribute to the proton gradient across the thylakoid.
Amino acid residues from the PSII membrane proteins D1, D2,
andCP43coordinate theassociationof theMn4CaO5cluster. The
removalof the fourelectrons fromwaterby theMn4CaO5cluster is
a four-step process. One electron is removed from water with
each photon-induced charge separation at the reaction center
and then passed via a redox-active tyrosine residue on the D1
protein (Yz) to fill the “hole” left in the PSII reaction center. Overall,
the removal of four electrons from two water molecules requires
the absorption of four photons by the reaction center that results
in the release of four H1 into the lumen. The electrons transferred
during the turnover of PSII are ultimately used to reduce two PQ
molecules to two PQH2 molecules, a process that also uses H1
from the stromal side of the thylakoid membrane.
Depending on the type of photosynthetic organism, at least
three extrinsic proteins are associatedwith the luminal face of the
complexwhereoxygenevolutionoccurs.Theseextrinsicproteins
are not required for oxygen production but do enhance it. These
proteins contribute to the stability of the Mn4CaO5 cluster,
facilitate the catalytic rate under physiologically relevant con-
ditions, andcontribute to theassemblyof thePSII enzyme.PsbO
is the only extrinsic protein conserved among cyanobacteria,
algae, and plants. This protein is also known as the manganese
stabilizing protein. Other extrinsic proteins include PsbP (plants
only), PsbQ (plants, cyanobacteria, and red algae), PsbU, and
PsbV (cyanobacteria and red algae). Thus, the complement of
extrinsic proteins associated with PSII varies among photosyn-
thetic organisms.
PSII Peripheral Antenna Complex or LHCII
Onemajor difference between plants and cyanobacteria is in the
structure of the peripheral protein/pigment complex that funnels
light to the photosynthetic core complex.
In cyanobacteria and red algae, the PSII accessory light-
harvesting system is composed of phycobilisomes, made up
of proteins and phycobilins. A central core anchors six rods
made up of phycobiliproteins and linker proteins that radiate
outwards. The entire phycobilisome structure is attached to the
stromal side of PSII and extends into the stroma.
In plants and green algae, the light-harvesting complex of PSII
sits in the thylakoid membrane. In vascular plants, it consists of
two layers: major, more abundant trimeric LHCII proteins and
minor, less abundant monomeric LHCII proteins. Based on
acrystal structure fromspinach (Spinacia oleracea), eachmono-
mer is made up of a single polypeptide chain, 14 chlorophylls,
and four carotenoids. In addition to the chlorophyll amolecules
found in the core complex, the outer LHCs also bind chlorophyll
b molecules and xanthophylls. The arrangement of the outer
layer varies with light and other conditions. Several additional
proteins are involved in the dynamic interaction between PSII
and LHCIIs. Under certain conditions LCHII complexes can
move away from PSII to decrease the amount of light funneled
into the PSII reaction center, providing flexibility to the photo-
synthetic system (see below). Furthermore, the ratio of light-
harvesting complexes to reaction centers varies with the light
intensity to which a plant is acclimated.
Q Cycle and Cytochrome b6f: Electron Transport from
Plastoquinol to Plastocyanin
Cytochrome b6f is a membrane-embedded complex that func-
tions as a dimer, with each dimer made up of eight subunits.
Some of the subunits are very similar to those in the cytochrome
bc1 complex that carries out electron transport inmitochondrial
respiration, and these complexes clearly share an evolutionary
origin. Theprotein subunits are cytochromeb6 andcytochrome f
proteins (hence the name of the complex), an iron-sulfur protein
also known as a Rieske protein (after its discoverer, John Rieske),
a 17-kD protein, and four shorter proteins. Unlike the related
complex in mitochondria, cytochrome b6f also binds chlorophyll
and b-carotene, as well as an unusual heme not found in the
mitochondrial complex.
Cytochrome b6f is sometimes referred to as plastoquinol-
plastocyanin oxidoreductase. Electrons from PSII are carried
by plastoquinol (PQH2) to cytochrome b6f where the Q-cycle
takes place. The Q-cycle (which also occurs in mitochondrial
electron transport) is a series of oxidation and reduction reactions
of that allows for maximal contribution to the proton gradient
and transfers electrons from a two-electron carrier (PQH2) to
a single-electron carrier (plastocyanin, a small protein). As elec-
trons are passed through cytochrome b6f, protons are passed
from the stroma to the lumen.
The Q cycle can be described as a series of steps. Reduced
PQH2 generated by PSII moves to the cytochrome b6f complex
where it is oxidized to PQ. During this initial oxidation, the release
of two protons into the thylakoid lumen results in one electron
6 The Plant Cell
transferred to PC, and the other electron to one of the two b-type
hemes borne by the complex and located on the opposite side of
themembrane.Another roundof oxidationoccurswhenasecond
PQH2 molecule delivers its electrons to cytochrome b6f. Again,
the release of one of its electrons results in the transfer of two
protons to the thylakoid lumen and one electron to plastocyanin.
Theother electron is transferred to another b-heme.Next, the two
reduced hemes cooperate to reduce a PQ into a PQH2, which is
then released from cytochrome b6f into the thylakoid membrane.
Thus, there is a recycled pool of PQ/PQH2 between PSII and
cytochrome b6f for the transfer of electrons between these two
complexes. The stoichiometry of the Q cycle is: two PQH2 enter,
two electrons are transferred to plastocyanin, four protons are
pumped into the lumen, and onePQH2 is regenerated; thus, each
molecule of PQH2 delivers four protons to the lumen and two
electrons to plastocyanin.
On the luminal side of the cytochrome b6f complex, electrons
are passed to themobile electron carrier plastocyanin, a copper-
containingenzyme. InChlamydomonasandcyanobacteria,butnot
plants,plastocyanincanbereplaced functionallybycytochromec6whencopper is scarce. Plastocyaninor (cytochromec6) transports
electrons from cytochrome b6f to PSI.
Structure and Function of PSI-LHCI
PSIuses light energy to transfer electrons fromthesolubleelectron
carrier plastocyanin on the luminal side of the thylakoidmembrane
to ferredoxinon thestromal side. It isamulti-protein,multi-pigment
complex that spans the thylakoid membrane and operationally
includes the core PSI complex and the peripheral antenna or light-
harvesting complex that is highly divergent.
Core PSI Complex
In terrestrialplants,PSI isacomplexof17proteinsubunitsandover
200 prosthetic groups, mainly, chlorophylls but also three Fe4S4
clusters and a few carotenoids and phylloquinones. The structure
is dominated by two large and related proteins, PsaA and PsaB.
Like in PSII, the heart of PSI is a pair of chlorophyll molecules,
one of which is excited and oxidized by light energy transduced
fromthesurroundingnetworkofpigmentmolecules.Thiselectron
moves through a chlorophyll to a phylloquinone and from there
through threeFe4S4 clusters andfinally to ferredoxin (Fd), a stable
electron acceptor. Ferredoxin is a soluble electron carrier that
binds reversibly to PSI and transfers electrons to NADP1 by the
action of the enzyme ferredoxin:NADP1 oxireductase. The electron
hole created when PSI absorbs a photon is filled by electrons
donated by the soluble electron transporter plastocyanin (or
cytochromec6). In cyanobacteria,PSI isusually foundasa trimer
but can also be present in tetramers or dimers, and it is thought
that this multimer arrangement increases the functional size of
the light-harvesting antenna for dim-light conditions.
PSI Peripheral Light-Harvesting or Antenna Complexes
As described for PSII, a key difference between plant, cyanobac-
terial, and algal PSI complexes is in the nature of the peripheral
light-harvesting complex. In plants and green algae, PSI is present
asamonomer that issurroundedbyanouterantennasystemmade
up of LHCI complexes that are embedded in the thylakoid mem-
brane and arranged in a half-moon shape. This crescent-shaped
arrangement ismadeupofasingle layerof fourproteins invascular
plants but a double layer of eight proteins in Chlamydomonas.
Bacterial PSI has not been purified with associated phycobili-
somes, but recently single-particle electron microscopy images
have been obtained and their energy transfer has been observed
by fluorescence spectroscopy.
Structure and Function of ATP Synthase
Proton translocation coupled to light-driven electron transfer
builds up the pH gradient (DpH) and electrical potential (DC)
across thylakoids. DpH and DC form the transthylakoid proton
motive force (pmf), which is the essential driving force for the
phosphorylation of ADP to produce ATP.
ATP synthase couples the synthesis of ATP fromADP andPi to
the movement of protons from inside the lumen (high concen-
tration; low pH) to the stroma (low concentration; high pH) across
the thylakoid membrane. The enzyme is also referred to as an
ATPase, as the reactioncanoccur inbothdirections (hydrolysis of
ATP linked to pumping protons against the gradient). The chlo-
roplast ATP synthase that carries out photophosphorylation is an
F-ATPase that is related to the mitochondrial ATPase and the
bacterial F-ATPase. The plant VH1-ATPase that is involved in
proton pumping across endomembranes is also an evolutionarily
related enzyme.
F-ATPases are large enzymes with more than 10 subunits that
assemble into two complexes. F0 is a membrane-embedded com-
plex that conducts protons and is made up of 10 to 15 copies of
subunit c, and one each of a, b, and b’ (in the older literature these
were referred to as subunits I to IV). F1 is a peripheral complex that
hydrolyzes/synthesizes ATP, is attached to F0 by a stalk, and is
made up of three copies each of thea andb subunits and one each
of the g, d, and e subunits. F1 is sometimes referred to as “coupling
factor 1” because when it is stripped off the membranes proton
efflux isuncoupled fromATPsynthesisduetothe lossof thecatalytic
complex of ATP synthase. Restoring F1 to stripped membranes
restores the coupling of proton efflux to ATP synthesis.
Paul Boyer and John Walker were awarded the 1997 Nobel
Prize in Chemistry “for their elucidation of the enzymatic mecha-
nism underlying the synthesis of adenosine triphosphate (ATP).”
Boyer laterwrote, “All enzymesarebeautiful, but theATPsynthase
is one of themost beautiful as well as one of themost unusual and
important,” and it is beautiful, particularly when seen in electron
micrographs. The enzyme uses a rotary catalytic mechanism
through which proton efflux provides energy for ATP synthesis.
Briefly, proton efflux through the F0 channel causes the a3b3
hexamer of F1 to rotate around the g subunit, and the conforma-
tional changes energize bond formation between ADP and Pi.
Thegandesubunitsworktogether toregulatethe levelofenzyme
activity, particularly in response to shifts between light and dark.
After a dark-to-light transition, the actions of PSII, cytochrome b6f,
and PSI cause protons to accumulate in the thylakoid lumen
and reduced ferredoxin and thioredoxin to be produced.
The chloroplast enzyme requires an acidified thylakoid lumen
November 2015 7
and reduced thioredoxin to be active, which prevents the
enzyme from running backward (hydrolyzing ATP) in the dark.
PATHWAYS OF ELECTRON TRANSPORT
There are at least three routes of photosynthetic electron trans-
port along the thylakoid membrane: (1) linear electron transport
that includes PSII, cytochrome b6f, and PSI; (2) cyclic electron
transport that includes PSI and cytochrome b6f; and (3) the
water-water cycle that uses PSII, cytochrome b6f, and PSI but
does not produce NADPH. The major pathway, linear electron
transport, is the onemost often taught, but alone it is inadequate
to meet the metabolic needs of the plant. The other pathways
serve to balance production of NADPH and ATP (both cyclic
electron transport and the water-water cycle produce a proton
motive force for ATP production but do not yield NADPH).
Furthermore, these alternative electron transport pathways
can provide a photoprotective function.
Linear Electron Transport Involves PSII and PSI
As described above, in LET, light energy transferred from the
LHCs to PSII and PSI oxidizes their reaction centers, driving the
linear transport of electrons from H2O to NADPH. The route of
linear electron transport can be summarized as:
H2O~PSII~PQ~Cyt b6 f~PC~PSI~Fd~NADP1~NADPH:
As a consequence of the Q-cycle, during LET the lumen gains
three protons for each electron that is transported: one proton
for each electron released by the oxidation of H2O at PSII and two
protons for each electron transferred through cytochrome b6f
to PSI. Therefore, linear electron transport has a proton/electron
(H1/e2) ratio of 3:1. The ratio of e2/NADPH is 2:1 (it requires two
electrons to reduceoneNADP1 toNADPH); thus, the ratio ofH1 to
NADPH is six. The stoichiometry of theATP synthase is such that it
takes 14 H1 to produce 3 ATPs (a ratio of H1/ATP of 4.67).
Consequently, the ATP/NADPH output ratio of LEF is 1.28 (which
equals 6/4.67). The pooled energy requirements for CO2 fixation in
theCalvin-Bensoncycle togetherwithphotorespirationandnitrate
assimilation inC3plantsare;1.43ATP/NADPH.Thus, LEF results
in a deficit of;0.15 ATP per NADPHproduced. This deficit can be
alleviated by cyclic electron transport, which passes electrons
cyclically throughPSIandcytochromeb6f leadingtothemovement
of protons into the lumen and subsequent ATP synthesis without
NADPH production.
Cyclic Electron Transport around PSI
Cycle electron transport (CET) uses all of the machinery of LET
except PSII. CET is a series of reactions in which electrons from PSI
reducePQtoPQH2,whichdeliversprotonstothelumenintheQcycle
ofcytochromeb6f.TheelectronsaretransferredtoPCandreturnedto
PSI to keepCETgoing; theelectronscycle. Protons released into the
thylakoid lumen contribute to the transthylakoid protonmotive force
(pmf). Therefore, CET contributes to the synthesis of ATP but not
NADPHandsoadjusts theATP/NADPHratioduringphotosynthesis.
There are two possible CET pathways. In both routes, PQH2
recycles electrons back to PSI through cytochrome b6f complex
and plastocyanin. The two routes of CET can be summarized as:
ð1ÞPSI~Fd~PQ~Cyt b6 f~PC~PSI ðdependent on PGR5=
PGRL1 in plantsÞ
ð2ÞPSI~Fd~NADP1~PQ~Cyt b6 f~PC~PSI ðdependent
on NDH complex Þ
The first route is sensitive to antimycin A and transports electrons
from PSI to Fd and then to PQ by ferredoxin plastoquinone re-
ductase. In Arabidopsis thaliana, the regulatory proteins PGR5
(PROTON-GRADIENT REGULATED5) and PGRL1 (PGR-LIKE1)
form a complex with PSI and mediate this antimycin-A sensitive
CET. This pathway appears essential for photosynthesis because
pgr5 and pgrl1 mutants have compromised photosynthesis. The
second CET route transfers electrons fromPSI to NADP1 through
Fd and then to PQ by NAD(P)H dehydrogenase complex (NDH
complex). The secondpathway is not essential for photosynthesis
because mutants deficient in NDH complex grow as well as the
wild-type plants in greenhouse conditions; however, the NDH-
dependent CET pathway may function under stress or becomes
essential when the first pathway is absent. Due to the difficulties to
measure CET accurately in vivo, controversies exist about the two
pathways of CET and how they are regulated.
The relative contribution of CET to photosynthesis depends on
themetabolicneedsof theplant.Thephotosyntheticneeds forATP
are higher in organisms or cells that concentrate CO2 such as
Chlamydomonas and C4 plants such as maize (Zea mays), so the
proportion of CET is greater than in C3 plants. Nevertheless, in C3
plants, CET seems to have a protective role in some conditions,
such as during the induction of photosynthesis or during low-CO2,
high-light, or drought conditions. Furthermore, by contributing to
the accumulation of H1 in the lumen, elevated CEF contributes to
photoprotective processes such as nonphotochemical quenching.
The Water-Water Cycle and Chlororespiration
The water-water cycle, also called the Mehler reaction, involves
the LET chain but uses oxygen instead of Fd as an electron
acceptor at PSI. Transferring electrons to O2 reduces it to super-
oxide (O2), which is subsequently reduced to H2O by superoxide
dismutase and ascorbate peroxidase. The water-water cycle
functions like a cycle because electrons are extracted from
H2O at PSII, transferred through PQ, cytochrome b6f, PC, and
PSI to reduce O2 and ultimately produce water. The water-water
cycle uses the linear transport chain and contributes to trans-
thylakoid pmf, but because the electron eventually goes to O2
instead of NADP1, it contributes to the production of ATP but not
NADPH. This pathway can be summarized as:
H2O~PSII~PQ~Cyt b6 f~PC~PSI~O2~H2O2~H2O:
The water-water cycle is thought to function for photoprotection.
During the light-induction phase (dark-adapted plants in light
condition), enzymes in the Calvin-Benson cycle initially are
8 The Plant Cell
not fully activated, so there is no consumption of NADPH for
CO2 fixation. Thewater-water cycle serves as a safety valve to
reduce the production of NADPHandbuild up a transthylakoid
proton motive force to trigger light dissipation through NPQ.
The water-water cycle has the disadvantage of generating
reactive oxygen species, which often lead to significant cel-
lular damage and must be quenched before damage occurs.
Chlororespiration is a light-independent reaction that transfers
electrons to oxygen and is also considered a mechanism to
alleviate excitation pressure on electron transport. Physiologically,
there are conditions in which the PQH2 pool becomes overly re-
duced, leavingnowhere for theelectronsproduced fromPSII togo.
Plastid terminal oxidase is a plastid enzyme, related to mitochon-
drialalternativeoxidase,whichoxidizesPQH2toPQbyreducingO2
to H2O. In other words, it draws electrons out from the electron
transport chain to prevent overreduction. Chlororespiration lowers
the excitation pressure on electron transport and likely reduces
overall oxidative stress. Chlororespiration links the redox state of
thePQpoolwith thatof thestroma,placingplastid terminal oxidase
at the crossroads of many metabolic processes.
In cyanobacteria, flavodiiron proteins (encoded by Flv genes)
are involved in photoprotection. Flv1 and Flv3 are necessary for
growth in fluctuating light conditions and function in aMehler-like
reaction that reduces O2 to H2O. Flv2 and Flv4 are thought to
protect PSII from damage by accepting electrons from the
electron transport chain. Furthermore, a thylakoid membrane-
associated terminal oxidase is also required to protect the cells
from damage during fluctuating light regimes.
DAMAGE AVOIDANCE AND REPAIR: ACCLIMATIONS TO
LIGHT STRESS
Photosynthetic organisms have to function in variable light
regimesandconditions. Theyare regularly subjected tovariation in
light intensity, light quality (relative amount of light of each wave-
length), and angle of light incidence.Someof these changes occur
over long time periods (seasons), others diurnally, and others on
the order of minutes, for example, the periodic sunflecks experi-
enced by leaves beneath a canopy. Variable light occurs against
a backdrop of variations in metabolism (e.g., energy demand and
nutrient availability) and environment (e.g., temperature and water
availability). Ashasbeendiscoveredbyscientists trying todevelop
artificial photosynthesis, it is rather amazing that such a high-
energy process can be maintained. Key to its success is the
ability to repair damage and to avoid it. Plants display numerous
strategies for both.
Photosynthetic organisms manage photon interception and
usage in the variable light environment to enable efficient pho-
tosynthesis and to minimize photodamage. As described pre-
viously, the rate of photosynthesis increases with light only at
relatively low light intensities, demonstrating that photosynthetic
organisms are regularly exposed to excess excitation energy.
Anytime when light energy exceeds the light saturation point,
plantsaresusceptible todamagedue to theoverreducedelectron
transport chain. When the system is light saturated, there is an
increased probability that excited chlorophyll will convert to the
excited triplet state (3Chl*),which iscapableof transferringenergy
tooxygen toproduce reactiveoxygenproducts.Therefore, plants
are exquisitely sensitive to the indicators of excess excitation
pressure, including a buildup of protons in the thylakoid lumen
and an overly reduced PQH2 pool; low lumen pH and high PQH2
levels each trigger regulatory responses to promote energy dis-
sipation. The timescale of responses can be extremely rapid (on
the order of seconds) and involve conformational or covalent
changes to the light harvesting machinery or occur more slowly
and involve changes in gene expression. Rapidly reversible re-
sponses to excess light are described as photoprotection,
whereas photodamage is reversed more slowly, and this differ-
ence is measureable.
Movements to Optimize Light Interception
Oneof themostdirectways tooptimize lightharvesting is tomoveor
to change orientation relative to sunlight. Single-celled organisms
can move toward light under light-limiting conditions to maximize
photosynthesis or away from excess light to prevent photodamage
inaprocesscalledphototaxis.Similarly,chloroplasts in leafcellscan
move to increase or decrease their interception of light through
accumulation or avoidance responses respectively. In many plant
species, leaf angle is sensitive to light, with the leaves rotating or
lowering at midday or other times when light incidence can exceed
photosynthetic capacity. In most cases, these movements are
mediated by light perception by photoreceptors such as photo-
tropins and phytochromes that are distinct from the pigments that
capture light forphotosynthesis,although there isalsoevidence that
the activity of PSII can affect leaf movement.
Acclimation via Stoichiometric Changes in Complex
Abundance
Arelativelyslowand long-termresponsetoprolongedlight intensity
changes is a change in the size of the light-harvesting systems, in
which their size is inversely proportionally to light intensity. These
changes arise through transcriptional and posttranscriptional con-
trolsofLHCgenesasa resultofsignaling fromthechloroplast to the
nucleus and involve both induction of expression of LHC genes in
low light and turnover of LHC proteins in high light. Similarly,
acclimation to high light involves an increase in the amount of
PSI, PSII, cytochrome b6f, and ATP synthase relative to the light
harvesting systems, a change that is reflected in an increased ratio
ofchlorophylla tochlorophyllb.Finally, lightspectralqualitiesaffect
the relative levels of PSII to PSI; specifically, in shaded canopies,
light is enriched for the longer wavelength light that is preferentially
absorbed by PSI, so the plant acclimates by increasing the abun-
dance of PSII. These acclimations can be quantified bymeasuring
the relative abundance of chlorophyll a to chlorophyll b and by
measuring the light saturation point.
Excess Light Energy Dissipation through
Nonphotochemical Quenching
Light energy absorbed by antenna or light-harvesting complexes
can have three fates. It can be processed for photochemistry,
November 2015 9
emitted via fluorescence, or dissipated byNPQ. The termNPQ
comes from the standard method for measuring photosyn-
thetic activities. By measuring the amount of chlorophyll
fluorescence under various conditions, it is possible to mea-
sure the amount of light that is quenched (i.e., not fluoresced)
due to photochemistry (termed photochemical quenching
[qP]) or NPQ.
NPQ can be divided into at least three different components
that can be distinguished by the relaxation kinetics of chlorophyll
fluorescence.Energy-dependentquenching (qE) isdependenton
the acidification of the lumen. It is the major and most rapid
component of NPQ in most algae and plants and relaxes within
seconds to minutes. State-transition quenching (qT) involves
movement of the mobile light-harvesting LHCII complex and
relaxes within tens of minutes. Photoinhibitory quenching (qI) is
caused by photoinhibition of photosynthesis and relaxes very
slowly in the range of hours. Each of these is described below.
Energy-Dependent Quenching (qE)
One consequence of excess light energy is a buildup of protons
in the thylakoid lumen that accumulate faster than they can be
processed by ATP synthase. Thylakoid lumen acidification is
sensed by special members of the LHC protein family that are
protonated on their luminal surface: PsbS in plants including
bryophytes, andLHCSR (light-harvestingcomplex stress related)
in Chlamydomonas and bryophytes. The acidic lumen also in-
duces the activity of an enzyme, violaxanthin deepoxidase,which
converts the carotenoid violaxanthin to zeaxanthin. This change
induces conformational changes in the PSII light-harvesting sys-
tem, causing intercepted light energy to be dissipated as heat
rather than transferred to the reaction center. The exact photo-
protective nature of this response continues to be debated and
may involve both transfer of energy from excited chlorophylls to
carotenoids in LHCII as well as charge transfer occurring in the
antenna complex that prevent light from reaching reaction center
chlorophylls.When the thylakoidpHreturns tonormal, zeaxanthin
is enzymatically convertedback toviolaxanthinand light energy is
again channeled into the reaction center.
Energy-dependent quenching is usually the dominant compo-
nentofNPQinplants.Thecontrol ofqEbyDpHallows inductionor
relaxation of qE within seconds in response to changing light
intensity to prevent excessive excitation of PSII centers under
high light or tomaximize light harvesting for photosynthesis in low
light. The acidification of the thylakoid lumen that results from
alternative electron transport, such as cyclic electron transport
and water-water cycle, promotes qE.
State Transition (qT)
Photosynthesis ismostefficientwhenPSI andPSII areoperating in
a balanced way, yet their antenna complexes have different com-
plementsofchlorophyllaandbandcarotenoids (thereforedifferent
absorption spectra), and light quality is variable, meaning that the
excitation energy transferred to PSI and PSII can be unbalanced.
One way to address this is through conformational changes in the
LHCII light-harvesting machinery that are referred to as a state
transition (qT). To some extent, this causes a channeling of light
energy fromLHCII to PSI, but it appears as though themajor effect
is to decrease light energy transfer to the reaction center of PSII.
State transition is thought to affect;20%of LHCII in Arabidopsis,
but as much as 80% of LHCII in Chlamydomonas.
Themolecular basis for state transition rests on the redox state
of the plastoquinone pool. Overexcited PSII leads to an accu-
mulation of reduced plastoquinone, PQH2.WhenPSII gets ahead
of PSI, the binding of reduced PQH2 to the cytochrome b6f
complex activates LHCII kinase; conversely, when PSI gets
ahead of PSII, the plastoquinone pool becomes oxidized and
LHCII kinase is inactivated. LHCII kinase was first identified in
Chlamydomonas and named Stt7, and later in Arabidopsis and
named STN7. Stt7 and STN7 are structurally and functionally
related and both are attached to thylakoid membranes. When
activated, they phosphorylate LHCII to induce state transition.
This reaction is reversible by the actionof a specificphosphatase.
Photoinhibition (qI)
Photoinhibition (qI) is light-induced reduction in photosynthetic
quantum yield that occurs as a consequence of photodamage
to the D1 protein of PSII (see below) or other slowly reversible
damage to the photosynthetic machinery. Light of any wave-
length can lead to photodamage, but plants can be particularly
sensitive to short wavelength blue or UV light hitting and dam-
aging the Mn4Ca cluster in the oxygen-evolving complex. This
damage slows the rate of electron flow from theOEC into the PSII
reaction center, which can lead to the production of reactive
oxygen species, which in turn decreases the rate of repair of the
D1 protein of the PSII reaction center. Photoinhibition can be
measured through fluorescence methods and is a slowly revers-
ible source of NPQ.
PSII Photodamage and Repair
While PSII requires light for its catalytic activity, this complex is
irreversibly damaged by light, more so than any of the other
complexes of the light-dependent reactions. PSII damage in-
creases with light intensity, but occurs to some extent at all light
intensities. The primary site of damage is the core D1 protein. To
recover activePSII complexes, thedamagedD1proteinmust be
recognized, proteolytically removed from the complex, and
replaced with a newly synthesized D1 protein. This involves
migration of the damaged PSII complex from the appressed
grana stacks to the unappressed regions, insertion of a newly
synthesized D1 polypeptide, and reassembly of the PSII com-
plex. To maintain a steady state level of functional PSII com-
plexes, photosynthetic organisms have dedicatedmachinery to
recognizeD1damageand repairPSII. It hasbeensuggested that
rather than being a defect, PSII’s sensitivity to photodamage
serves as a safety valve to protect PSI from damage.
Sensitivity of Photosynthesis to Heat, Drought, and Other
Stress
As described above, many short-term and long-term acclima-
tions enable photosynthetic processes to be maintained even as
10 The Plant Cell
light intensity changes. Essentially any kind of stress, ranging
fromnutrient stress,drought stress, temperaturestress, andeven
pathogen attack, can affect carbon fixation, which in turn affects
the flow of electrons through the photosystems as well as the
cycling of ATP/ADP, NADPH/NADP1, and PQ/PQH2. Changes in
DpHand redox state of thePQpool are important in regulating the
core photosynthetic processes as well as pathways to confer
protection from photooxidative damage.
However, there are additional ways that some stresses can
affect photosynthesis. For example, high temperatures increase
thefluidityof the thylakoidmembrane,which inturnaffectselectron
transport, proton efflux through ATP synthase, and ionmovement
across thylakoid membranes. One response to increased thyla-
koid leakiness is an increase in CET to maintain proton motive
force. As another example, stomatal closure in response to water
deficitdecreases theuptakeofCO2,meaning that thepoolsofATP
and NADPH accumulate, resulting in an overreduced electron
transport chain and the accumulation of protons in the thylakoid
lumen. This increase in transthylakoidprotonmotive force induces
an increase in energy-dependent quenching, which helps to dis-
sipateexcess lightenergyandprevent light-induceddamage.Cold
temperatures slowmetabolic processes but do not affect the rate
of light interception, and cold temperatures are frequently asso-
ciated with an increase in photooxidative damage. One of the big
ongoing research areas is to integrate what we know about the
light-dependent reactions under optimal conditions with the
whole-plant physiological changes that occur with stress.
Regulation and Retrograde Signaling
Many of the biochemical shifts and responses described above
occur autonomously in plastids and can be measured in isolated
chloroplasts. However, other photosynthetic acclimations and
stress responses involve changes in nuclear gene expression.
Not only are most of the chloroplast-localized proteins encoded
by genes in the nucleus, but the chloroplast operates within the
larger context of the cell and the rest of the cell’s complement of
nuclear-encoded proteins. Thus, there is a clear need for in-
tegration between the functions of the plastid, the nucleus, the
cytosol, and the mitochondrion.
Information flows from chloroplast to nucleus. This has been
shown in several ways, for example, through the observation that
plastid ribosomal protein mutants, which have abnormal plastid
activities, also show a downregulation of nuclear-encoded pho-
tosynthetic genes, an observation that reveals that information
about plastid functions is signaled to the nucleus. Along with
similar signals from the mitochondria, these are known as retro-
grade signals; signals from the nucleus to the organelles are
known as anterograde signals.
Other than their existence, the nature of retrograde signals
remains uncertain. A current view is that multiple signals convey
different types of information, as indicated by different tran-
scriptional responses. Furthermore, distinct signals are thought
to be involved in nuclear processes during chloroplast devel-
opment (biogenic controls) versus those involved in fine-tuning
mature chloroplast functions (operational controls) in response
to changing conditions.
Several different retrograde signals have been identified and
there isconflictingevidence for the relative importanceofdifferent
types of signals. One type of signal was first identified from a
genetic screen for an uncoupling of the nuclear and plastid ge-
nomes; the identifiedmutants are knownasgenomeuncoupled or
gun mutants, and several affect heme biosynthesis. Because
tetrapyrroles, including chlorophyll or heme or their biosynthetic
intermediates, can cause light-mediated damage, their levels are
necessarily subject to tight regulation. However, the nature of the
heme-derived signal, whether an intermediate or an indication of
flux rate, remains obscure. Another signal identified through ge-
netic approaches is PAP (3#-phosphoadenosine 5#-phosphate).PAP is synthesized in plastids andPAP levels increase inwild-type
plants exposed todrought or high light intensity. PAP is an inhibitor
of exoribonuleases and is thought to affect RNA metabolism and,
therefore, gene expression. Finally, there is evidence to support
various reactive oxygen species and even a chloroplast-envelope
bound transcription factor (which is cleaved off the plastid and
translocated to the nucleus under stress) as having roles as
retrograde signals. The nature and roles of retrograde signals
continue to be very actively investigated.
MONITORING LIGHT REACTIONS
Insights into the light-dependent reactions can be obtained
from gas exchange studies or analysis of the fluorescence/
absorbance properties of the reaction center chlorophyll and
other pigments.
Gas Exchange Analysis of Photosynthesis
Photosynthesis involves the consumption of CO2 and the pro-
duction ofO2, and achange in the concentration of these gasses
provides a convenient way to measure photosynthesis. The
measurements can be made in a chamber clamped to a leaf
or by placing an entire plant into a sealed chamber. CO2 absorbs
infrared light so its concentration is relatively easy to measure
spectroscopically, whereas oxygen can be measured using oxy-
genelectrodes.CO2measurementsarequite informativeabout the
finalprocessesofphotosynthesis, carbonfixation,butarenot ideal
for the study of light-dependent reactions as there are so many
additional steps and variables (e.g., CO2 concentration, stomatal
conductance, Rubisco activity, etc.) between carbon fixation
and the reactions that take place in the thylakoid membranes.
Furthermore, both O2 and CO2 levels are affected by mitochon-
drial and peroxisomal metabolism, so net gas exchange mea-
surements represent sums of many processes.
Spectroscopic Measurements of Light-Dependent
Reactions
Several spectroscopic measurements are available to monitor
light reactions of photosynthesis in vivo using light-adapted,
intact leaves. PSII activity can be measured by chlorophyll
fluorescence, PSI activity can be measured by the absorbance
change at 810 to;830nm, and the transthylakoid protonmotive
November 2015 11
force and proton flux can be measured by electrochromic shift
(ECS). More details are provided below.
Chlorophyll Fluorescence Measurement of PSII Activity
When a chlorophyll amolecule absorbs light, it is excited from
its ground state to its singlet excited state (Chl*), which can
return to the ground state via three processes: (1) the excita-
tion energy can be transferred to reaction centers to drive
photosynthesis through photochemistry (qP); (2) the energy
can be released as heat through NPQ; (3) the energy can be
reemitted as chlorophyll fluorescence. At room temperature,
chlorophyll fluorescence mainly originates from PSII, so chlo-
rophyll fluorescence conveys quantitative information about
PSII photochemistry.
Fluorescence yield is usually low (0.6 to 3% of total light
absorbed); however, because the fluorescence emission peaks
at longer wavelength than absorbed light, it can be measured by
illuminating a leaf with light of a defined wavelength and mea-
suring theamountof fluorescenceemissionat longerwavelength.
In order to measure fluorescence in daylight, a modulated mea-
suring system can be used, in which the light used to induce
fluorescence (known asmeasuring light) is modulated (switching
on andoff at high frequency) and thedetector is tuned todetect at
the same frequency.
First, a baseline level of fluorescence (Fo) must bemeasured in
dark-adapted leaves (in which the photosynthetic enzymes and
intermediates and nonphotochemical adaptations are all as-
sumed to be negligible), using light intensity that is too low to
initiate photochemistry. Next, the maximum yield of fluores-
cence can bemeasured. A short saturating pulse of light causes
all of the photosynthetic reaction centers to give up an electron
and become reduced, transiently preventing further photo-
chemistry; the reaction centers in this state are described as
closed. Therefore, this pulse is followed by the maximal level of
fluorescence, denoted Fm. The difference in fluorescence signal
between Fm and Fo is the variable range of fluorescence, denoted
Fv, and the ratio of Fv to Fm indicates the maximum quantum
efficiency of PSII.
Next, actinic light (light sufficient to support photosynthesis)
canbeswitchedon.Again, a rapid fluorescence response follows
(due to the closing of reaction centers), but the fluorescence
decays as the photosynthetic enzymes are activated and pho-
tochemical and nonphotochemical energy dissipation processes
ramp up. This quenching of fluorescence is due to a combination
of nonphotochemical and photochemical quenching.
Todistinguish between photochemical and nonphotochemical
quenching of chlorophyll fluorescence, a brief (#1 s) flash pulse is
used to transiently saturateandclose (reduce) all thePSII reaction
centers, resulting in zerophotochemical quenching. It is assumed
that the flash is short enoughso that no (or a negligible) increase in
the NPQ occurs and no long-term change of photosynthesis is
induced. The flash causes the chlorophyll fluorescence to reach
a maximum value (Fm#, the prime denotes the fluorescence in
light-adapted leaves) which is attained in the absence of any
photochemical quenching.
The difference between Fm (dark-adapted leaves) and Fm#(light-adapted leaves) is ameasure of NPQ, and the rate at which
maximal fluorescence is restored in the dark varies by the type of
NPQ. For example, qE reverses more quickly than qI.
The difference between Fm# (maximal fluorescence in light-
adapted leaveswhen reaction centers are closed) and the steady
state yield of fluorescence (F#, also written as Fs#) indicates the
extentof photochemistry occurring in the light (which ismeasured
here as photochemical quenching). In other words, Fm#2 F# is anindicationof theperformanceofPSII, andwhennormalized toFm#is described as the PSII operating efficiency or FPSII.
Under optimal conditions,FPSII is usually linearly proportional
to carbon fixation rate measured by gas exchange. However,
under stressful conditions (e.g., light, drought, and temperature
stress), this linear relationship often does not hold and FPSII
overestimates the rate of carbon fixation, because stress-
induced alternative electron transports (e.g., water-water cycle;
described above) can contribute to the total electron transport
throughPSII but theproduct isnotused for carbonfixation.Thus,
chlorophyll fluorescence in combination with gas exchange is
a powerful and accurate way to monitor photosynthesis under
diverse conditions.
P700 Measurement for PSI Activity
PSI photochemistry is initiated by light energy transfer from
antenna pigments associated with PSI to its reaction center
chlorophyll (P700), which absorbs light and goes to the excited
state (Chl*). The electron is then transferred from the excited
state Chl* to a downstream primary electron acceptor, eventually
producing oxidized P700, which is denoted as P7001. P7001
absorbs 810-nm light (peak at 810 to ;840 nm), while P700
does not, so the oxidation status of P700 can be determined by
absorbance at 810 nm.
To measure the percentage of oxidized P700, the maximum
oxidizable P700 pool is attained by applying a saturation flash
along with far-red light (which preferentially excites PSI), which
together ensures the complete oxidation of the P700 pool. The
P700 oxidation ratio is determined by the ratio of P7001 induced
by actinic light to the maximum amount of P7001 (induced by far
red and flash). Upon turning off the light, the reduction rate of
P7001 can be measured.
An increase in the relativepoolof reducedP700can indicate the
occurrence ofCEF. The rate ofCEF is a key acclimation to several
environmental variables; for example, plants inwhichCO2fixation
is limited tend to showan increase inCEF,which canbeobserved
by PSI activity measurements.
Electrochromic Shift to Monitor Transthylakoid Proton
Motive Force
Charge movements across the thylakoid membrane affect the
transthylakoid electric field, which in turn affects carotenoid ab-
sorbance. Thepeakchange incarotenoidabsorbanceat518nm is
knownastheECS,and itcanbeusedtomonitor thepmfacross the
membrane. The full amplitude of the ECS absorbance is propor-
tional to the total light-induced pmf, and following a dark interval,
theECSdecay rate reflectshowfastprotonsmoveoutof the lumen
through ATP synthase. ECS provides a noninvasive, direct, in situ
measurement of the thylakoid energization and deenergization,
12 The Plant Cell
transthylakoid pmf, and proton conductance of the thylakoid
membrane. Furthermore, by comparing the full amplitude of
ECS absorbance (which is proportional to pmf) in the absence
and presence of PSII inhibitors, the contribution of cyclic electron
transport to pmf can be estimated.
ECS measurements can be extended to measure the two
components of pmf. During a short dark interval (;500 ms),
ECS is dominant and measurement at 518 nm is sufficient to
measure the transthylakoid pmf; however, during longer periods,
other components that occur on the minutes timescale, like zeax-
anthin formation (absorbance change peak at 505 nm) and light
scattering (absorbance change peak at 535 nm), overlap and
interfere with the ECS. To circumvent this, multiwavelength
measurements and deconvolution are necessary to subtract
the effects of zeaxanthin and light scattering from that of the
ECS. Multiwavelength measurements can be deconvoluted so
that ECS can bemonitored during a longer dark interval (25 s) to
separate the two components of the transthylakoid pmf, DpH,
and DC.
Fluorescence Imaging
By combining light emitting diodes and sensitive digital cameras,
it is possible to incorporate spatial dimensions into fluorescent
measurements and represent these measurements as images.
These methods can be particularly useful in understanding how
photosynthetic parameters vary across a leaf. Furthermore,
although they lack the precision of fine-scale imaging systems,
it is possible to engineer systems capable of imaging fluorescence
over larger scales such as fields, which can help to automate the
screening process when breeding for enhanced resilience to
stress. In some cases, these systems rely on solar-induced fluo-
rescence,which isnotascontrolledaspulseamplitude-modulated
measurements and so more difficult to interpret.
OPTIMIZING AND IMPROVING PHOTOSYNTHESIS
Natural selection produced plants that survive in their natural
environments. We depend on plants or algae that produce high
yields of specific products in what may be very unnatural envi-
ronments. Given that plants in natural environments are often
growth-limited by water or nutrients rather than sunlight, it is
possible that they are not genetically optimized for high photo-
synthetic capacity. Investigations into improving light-dependent
reactions can be summarized as: (1) harvesting otherwavelengths
of light more efficiently, (2) decreasing shading, and (3) minimizing
photooxidativedamageandaccelerating repair.At thispoint,most
of these explorations are quite preliminary or theoretical, but
nevertheless interesting.
Harvest Light More Efficiently
Plants use only a narrow range of light wavelengths for photosyn-
thesis (400 to;700 nm), and green light is only weakly absorbed
(reflected/transmitted green light gives leaves their color).
Engineering plants to express phycoerythrin, a phycobiliprotein
from red algae that absorbs green light, might increase the
capture and flow of photons from the light-harvesting complex
to the reaction center chlorophyll, provided that the pigment
could be properly anchored and oriented. Another possibility
could be to introduce bacteriochlorophylls that absorb longer
wavelengths on the order of 1000 nm. However, using this
longer wavelength light would require extensive remodeling of
the photosynthetic apparatus, as it does not have enough
energy to drive reactions in the current system of oxygenic
photosynthesis.
Truncate Light-Harvesting Antenna Systems
In any photosynthetic system, shading occurs. Is it possible to
decreaseshading formoreeffective lightharvesting?Perhaps the
easiest system in which to apply this idea is a cell culture system
like those grown for the production of biofuels. Cells on the
surface may intercept too much light and need to dissipate the
energy, while at the same time those on the interior of the culture
vessel can be light-limited. In cyanobacteria cultures with trun-
cated light-harvesting antenna, overall levels of photosynthesis
were enhanced due tomore light being available to shaded cells.
Can this strategy be applied in multicellular plants? We know
that plants acclimate to a light gradient in several ways, for
example, by differences in the size of the antenna complexes
relative to the reactioncenters inupperversus lower leaves (which
is easily measureable by a change in the ratio of chlorophyll a to
chlorophyll b). It might be possible to engineer plants with sig-
nificantly decreased light absorption capacity in their upper
leaves (through both molecular and anatomical changes such
as amore vertical orientation) and similarly increased capacity for
light absorption in the lower leaves. Could photosynthetic ca-
pacity be increased by truncating antenna complexes in the
complex three-dimensional leaf canopy?
Minimizing Damage and Accelerating Repair
Currently the most feasible strategy for optimizing photosyn-
thesis may be to decrease photooxidative damage. When
excess excitation energy is not dissipated efficiently, reactive
oxygen species can be produced, and the damage caused can
be metabolically costly to repair. Several studies suggest that
damage can be minimized by engineering plants to be more
sensitive to excess excitation energy and to more effectively
control it through enhancing the xanthophyll cycle or by
reducing the PQH2 pool via CEF or chlororespiration. For
example, a protein present in cyanobacteria and bryophytes,
LHCSR, senses excess light energy and triggers NPQ. It has
been suggested that introducing this protein into vascular
plants could similarly decrease photooxidative damage and
so enhance photosynthesis. Similarly, photosynthesis could
be enhanced by increasing the rate of replacement of the D1
protein of PSII. Further explorations of these strategies will
require an analysis of the metabolic costs associated with
enhanced photoprotection and to what extent they decrease
quantum efficiency under optimal conditions.
November 2015 13
ARTIFICIAL PHOTOSYNTHESIS
The energy that reaches the earth from the sun in an hour is
equivalent to “all the energy humankind currently uses in a year”
(Barber and Tran, 2013). An early proponent of harnessing this
energywas the Italian scientistGiacomoCiamician (1912),whose
vision for a smokestack-free future sounds remarkably contem-
porary, “And if in a distant future the supply of coal becomes
completely exhausted, civilization will not be checked by that, for
life and civilization will continue as long as the sun shines!” More
than 100 years later, chemists and engineers are still working to
develop an affordable, effective, and efficient way to fuel civili-
zation with energy obtained from sunlight.
Artificial Photosynthesis for Chemical Energy Production
Thecostofconvertingsunlight toelectricalenergy isbecomingmore
affordablewith thedevelopmentof improvedphotovoltaic systems.
However, the limitationof thesesystemsis that thecurrentproduced
is difficult to store, and they are most effective when the energy is
used immediately; in other words, your solar-powered light is less
efficient after dark, which is when you need it most.
An alternative approach is to use solar energy to produce
chemical energy, which can be stored more easily and used
when and where it is most needed. This process is often de-
scribed as artificial photosynthesis. As in biological photosyn-
thesis, the process can be broken down into four steps: (1) light
harvesting, (2) using light energy to separate charge across
a membrane, (3) the oxidative step, using that charge separation
to oxidize water, and (4) the reductive step, using that charge
separation to reduce a substrate (such asH1 to H2 gas, or CO2 to
formic acid and other organic compounds). Artificial photosyn-
thetic systems can be entirely synthetic or can incorporate prop-
erties of living cells and synthetic biology.
One challenge is to develop semiconductingmaterials that can
support charge separation and the catalysis ofwater splitting and
hydrogen generation. Some materials can do this without addi-
tional electrocatalysts, or these can be provided separately in
a hybrid system.Difficulties include findingmaterials that are able
to capture sufficient energy using a broad spectrum of light (as
opposed to just themost highly energetic UV light), materials that
are abundant enough in the earth’s crust to be feasible for large-
scale production, andmaterials that are durable and persistent in
the functioning device. Some of the ideas being explored are
directly inspiredbyphotosynthesis, for example, thecoupling two
light-harvesting devises in series, one to drive water splitting and
one to drive hydrogen reduction, and the use ofMnCacomplexes
as catalysts for the water-splitting reaction.
Another approach that can be described as semisynthetic in-
corporates enzymes or even living cells into an energy production
system. As examples, hydrogenase enzymes can increase the
efficiency of hydrogen productionwhen incorporated into synthetic
systems, natural chromophores can enhance light harvesting in
synthetic systems, and hybrid systems composed of catalysts
introduced into cell culture systems can increase the efficiencies
by which light is converted to biofuels.
Alternatively, a synthetic biology approach can be used, in
which the properties and functions of living cells are altered so
they make desired products more efficiently. As an example,
Chlamydomonas can be engineered to more efficiently produce
H2 for use as fuel, and bacteria or algae or diatoms can be
metabolically engineered to increase their production yield of
useful biofuels. Regardless of the approach, the Second Law of
Thermodynamics must be followed. In any energy utilization
process, it is imperative that getting rid of entropic waste does
not exceed the value of the captured light energy. Currently none
of these strategies are highly efficient, but the goal of cheap and
abundant energy is propelling this research at a fast pace.
Rhodopsin-Based Phototropy
Finally, energy from the sun can be harnessed using systems
based on rhodopsins. It is important to note that rhodopsins are
entirely unrelated to the photosystems we have been describing
that arebasedonchlorophyll, but theyare introducedhereas they
show promise as energy-harnessing systems. Rhodopsins are
light receptorsmade up of an opsin protein and retinal, a pigment
related to vitamin A. Rhodopsins act as light sensors (they are
the photoreceptors found in animal eyes), but in prokaryotes they
can harness light energy through the process of phototropy.
Rhodopsins do not participate in electron transfer reactions
and therefore do not permit full photosynthesis. Bacteriorhodop-
sins use light energy to pumpprotons across amembrane, which
can be used for secondary transport or ATP synthesis; other
systems pump chloride. Rhodopsin systems have spread widely
throughout prokaryotes throughgene transfer andare genetically
simpler than chlorophyll-based photosynthetic systems, requir-
ing only a few genes to produce retinal and opsin. This genetic
simplicity makes them attractive targets for synthetic biology
approaches. However, rhodopsin-based systems use light less
efficiently than chlorophyll-based systems, due to both a nar-
rower light action spectrum and a lower efficiency of coupling
protons to chemical energy. Nevertheless, several efforts are
underway to harness them as light-energized pumps.
PHOTOSYMBIOSIS: PHOTOSYNTHETIC FUNGI AND
ANIMALS
Not all photosynthetic organisms are plants, algae, or bacteria.
We conclude this lesson by describing members of fungal and
animal kingdoms that have acquired the ability to live symbiot-
ically with photosynthetic organisms in a process that has been
described as photosymbiosis. A few are oddities, but others are
important contributors to Earth’s ecosystems; in fact, it is esti-
mated that 50%of theocean’sphotosynthetic output is a result of
photosymbiosis.
Lichen: Fungus1 Algae or Cyanobacteria
Lichen are widespread and provide a valuable food source for
many animals. Lichen are symbiotic organisms that consist of
a fungus (themycobiont) and an extracellular photobiont that can
becyanobacteria (in10%of lichens) orgreenalgae (in90%); a few
are tripartate and include both an algae and cyanobacterium.
14 The Plant Cell
The symbiosis can persist through vegetative propagation or be
reestablished following spore formation. Nearly 20,000 lichen
species are known. They are classified by the name of the
dominant fungal partner and have been described as the sym-
biotic phenotypeof the fungus. In thismutualistic association, the
photobiont provides reducedcarbonand themycobiont provides
water, nutrients, and a sheltered environment for the photobiont.
The whole is greater than the sum of the parts and lichen are able
to live in a wide range of environments and grow all across the
world on an amazing variety of substrates.
Photosynthetic Animals
Many types of marine invertebrates are able to serve as hosts
for photosynthetic algae, although it is not always clear if this
relationship is truly beneficial to both partners. Many marine
plankton that form the foundation of the ocean’s food web are
protists from the taxa Rhizaria, Alveolates, and Stramenopiles
that harbor microalgal symbionts.
Some of the more familiar photosynthetic animals are the
photosynthetic corals, which are simple animals in the phylum
Cnidaria. Some corals form calcium carbonate structures,
including huge reefs that provide important ecosystems for
marine organisms. The ability of corals to produce these vast
carbon-based structures is largely dependent on their mutual-
istic relations with a type of algae known as dinoflagellates or
zooplankton. These symbioses can be passed on maternally or
become established in vegetative tissues. Warming and acid-
ifying oceans can cause the symbiont to die or be expelled from
the host, resulting in bleaching and death of the coral. A recent
study indicates thatwe are currently in themidst of a global coral
die-off as a consequence of warming oceans and El Nino
weather events.
A tidal acoel flatworm, Symsagittifera roscoffensis (formerly
known as Convoluta roscoffensis), is providing a good model for
investigating photosymbiosis, in part because it lacks coral’s
calcified structures. Larvalworms ingest greenalgae (Tetraselmis
convolutae) a few days after hatching. Once ingested, the algae
divide mitotically and persist extracellularly within the worm’s
body. Adults can host 40,000 algal cells and appear bright green
(an early researcher describe them as “plant-animals”). These
organisms are providing insights not only into the biochemistry of
photosymbiosis but also how the symbiont affects the host’s
behavior; like other photosynthetic organisms, these worms
exhibit positive phototactic behavior.
Most of the known algal-animal symbioses involve inver-
tebrates, but there is one example that involves a vertebrate.
Eggs of the spotted salamander Ambystoma maculatum appear
green due to the presence of a single-celled algaOophila amblys-
tomatis (oophila means egg-loving). The algae are found only in
association with the salamanders and are thought to benefit
from the nitrogenous waste produced by the developing
embryo. The animal is thought to benefit from the oxygen
released as a consequence of photosynthesis. Although this
was thought to be an extracellular symbiosis, algae have
recently been identified living transiently within the animal
cells, the first occurrence of an intracellular endosymbiosis in
a vertebrate host.
Kleptoplastic Sea Slugs
In each of the above cases, the animals host intact unicellular
algae, in what is clearly a symbiosis of two organisms. The sea
slug Elysia chlorotica reveals a different approach. These
animals eat algae and then maintain the algal chloroplasts in
an intact and functional state within their digestive tissues, in
a process known as kleptoplasty (plastid-stealing). The stolen
plastids have been shown to remain photosynthetically active
for several months after removal from the algal cell. Although it
was once proposed that some of the algal genes had migrated
to the sea slug nucleus, this idea has largely been discredited,
so this interestingsymbiosis raises thequestionofhowaplastid
stays viable when it cannot be restored by gene products from
the nucleus.
Although salamanders and sea slugs do not contribute
much to the global energy economy, they provide novel re-
search models and novel opportunities to fuel imaginations
and ignite interest in the vitally important, life-sustaining
reactions of photosynthesis.
SUMMARY AND ONGOING RESEARCH
The question of how plants convert light energy into chemical
energy has fascinated scientists since it was first recognized.
Todaywehaveagoodunderstanding of the chemical processes
as well as the structure of the molecular machines that carry
them out. Great strides in obtaining structural information of the
photosynthetic complexes has led to atomic-level understand-
ing of the core reactions, which has been augmented by com-
parative studies of photosynthesis in organisms that diverged
from each other millions of years ago. New approaches to the
study of photosynthesis are refining our understanding of the
extremely rapid events that occur as light is captured, such as
quantum coherent energy transfer that excites physicists and
can contribute to more efficient light harvesting efficiency in
solar energy cells.
As we continue to delve more deeply into defining the core
reactions that harvest light, we also are expanding our un-
derstanding of how they respond to fluxes of light and me-
tabolism. In multicellular organisms, we have the additional
questions of how developmental processes and intercellular
signals and fluxes influence and are influenced by the light-
dependent reactions of photosynthesis. Genetic approaches
through mutants and new approaches including metabolo-
mics and systems approaches are increasingly important in
building a complete understanding that integrates processes
that span broad scales of time and space.
Opportunities are being explored for improvements in
photosynthetic efficiency, to enhance production from plants
and cultured cells. Opportunities are also being explored
for the development of alternative energy sources such as
biofuels from single-celled algae or cyanobacteria, or bio-
inspired artificial photosynthesis. Perhaps the biggest chal-
lenge for the 21st century will be to learn from plants how to
harness and use the abundant energy that the sun provides to
the Earth.
November 2015 15
Ru Zhang
Department of Plant Biology
Carnegie Institute for Science
Johnna Roose
Louisiana State University
Mary Williams
Features Editor, The Plant Cell
c/o Laboratory of Plant Physiology and Biophysics
University of Glasgow
ORCID ID: 0000-0003-4447-7815
RECOMMENDED READING
(This is a representative list of sources to help the reader access
ahugebodyof literature.Weapologize inadvance to thosewhose
work is not included.)
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