Lighting the way: Compelling open questions 
in photosynthesis research
Nancy A. Eckardt,1,*,† Yagut Allahverdiyeva,2 Clarisa E. Alvarez,3 Claudia Büchel,4 Adrien Burlacot,5,6 Tanai Cardona,7,8
Emma Chaloner,7,8 Benjamin D. Engel,9 Arthur R. Grossman,5,6 Dvir Harris,10 Nicolas Herrmann,4 Michael Hodges,11
Jan Kern,12 Tom Dongmin Kim,7,8 Veronica G. Maurino,13 Conrad W. Mullineaux,7 Henna Mustila,2 Lauri Nikkanen,2
Gabriela Schlau-Cohen,10 Marcos A. Tronconi,3 Wojciech Wietrzynski,9 Vittal K. Yachandra,12 Junko Yano12
1The Plant Cell, American Society of Plant Biologists, USA
2Molecular Plant Biology Unit, Department of Life Technologies, University of Turku, 20014 Turku, Finland
3Centro de Estudios Fotosintéticos y Bioquímicos (CEFOBI-CONICET), Facultad de Ciencias Bioquímicas y Farmacuticas, University of Rosario, Suipacha 570, 2000 
Rosario, Argentina
4Institute of Molecular Biosciences, Goethe University Frankfurt, 60438 Frankfurt, Germany
5Division of Bioscience and Engineering, Carnegie Institution for Science, 260 Panama Street, Stanford, CA 94305, USA
6Department of Biology, Stanford University, Stanford, CA 94305, USA
7School of Biological and Behavioural Sciences, Queen Mary University of London, Mile End Road, London E1 4NS, UK
8Department of Life Sciences, Imperial College London, London SW7 2AZ, UK
9Biozentrum, University of Basel, Sptialstrasse 41, 4056 Basel, Switzerland
10Department of Chemistry, Massachusetts Institute of Technology, Massachusetts Ave, Cambridge, MA 02139, USA
11Université Paris-Saclay, CNRS, INRAE, Université d’Evry, Université de Paris Cité, Institute of Plant Sciences Paris-Saclay (IPS2), 91190 Gif-sur-Yvette, France
12Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
13Molecular Plant Physiology, Institute for Cellular and Molecular Botany (IZMB), University of Bonn, Kirschallee 1, 53115 Bonn, Germany
*Author for correspondence: neckardt@aspb.org.
†Authors are listed alphabetically (except for the lead author/coordinating editor).
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for 
Authors (https://academic.oup.com/plcell) is: Nancy A. Eckardt (neckardt@aspb.org).
Abstract
Photosynthesis—the conversion of energy from sunlight into chemical energy—is essential for life on Earth. Yet there is much we do not 
understand about photosynthetic energy conversion on a fundamental level: how it evolved and the extent of its diversity, its dynamics, 
and all the components and connections involved in its regulation. In this commentary, researchers working on fundamental aspects of 
photosynthesis including the light-dependent reactions, photorespiration, and C4 photosynthetic metabolism pose and discuss what 
they view as the most compelling open questions in their areas of research.
Received January 23, 2024. Accepted July 15, 2024 
© The Author(s) 2024. Published by Oxford University Press on behalf of American Society of Plant Biologists. 
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which per-
mits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.
Introduction
(Written by Nancy A. Eckardt)
In the garden on a September morning, I was, once again, aston-
ished to find that a tiny seed I planted a few months ago had pro-
duced large stems, leaves, and—seemingly daily—dozens of 
enormous fruit (also known as zucchini), literally out of thin air. 
This plant took carbon as CO2 directly out of the air and turned 
it into organic matter, and along the way, split water molecules 
to produce oxygen and stored energy, in one of the most remark-
able and essential life processes: photosynthesis.
All oxygenic photosynthetic organisms have two photosystems 
(PS) known as PSI and PSII, which are embedded in thylakoid 
membranes and carry out light-driven electron transport. PSII ab-
sorbs light energy of wavelengths <680 nm to oxidize water into 
molecular oxygen (the “water-splitting” reaction), which contrib-
utes to the proton gradient needed to convert ADP to ATP, whereas 
PSI absorbs energy of longer wavelengths to generate high-energy 
electrons used to convert NADP+ to NADPH. All PSs have an array 
of light-harvesting antennae and mechanisms to dissipate excess 
energy under fluctuating light conditions. Many of the core 
reactions and features of the PSs are highly conserved among pho-
tosynthetic organisms. Research on the structure and function of 
the core components of the photosystems and thylakoid mem-
branes often uses phototrophic microorganisms for the ease of 
maintaining cultures and isolating or purifying components. 
These include the green alga Chlamydomonas (Chlamydomonas 
reinhardtii), diatoms such as Phaeodactylum and Thalassiosira, and 
cyanobacteria such as Synechocystis and Synechococcus. With mi-
croorganisms, it is also possible to study in vivo and whole organ-
ism properties of photosynthesis without some of the many 
confounding processes present in land plants, such as stomatal 
conductance of CO2, water relations, and carbohydrate allocation 
(source–sink) dynamics.
A lot of photosynthesis research today is directed toward im-
proving photosynthesis to create more climate-resilient crops 
and enhance crop yields in the face of global climate change 
(see the companion article in this issue by Croce et al. 2024). And 
yet, despite the long history of photosynthesis research (see e.g. 
Nelson and Junge 2015; Stirbet et al. 2020; Sharkey 2022) and sig-
nificant advances we have seen in recent years (e.g. Croce and van 
Amerongen 2020; He et al. 2020; Perera-Castro and Flexas 2020; 
The Plant Cell, 2024, 36, 3914–3943 
https://doi.org/10.1093/plcell/koae203 
Advance access publication 22 July 2024 
Commentary
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Bhowmick et al. 2023; Greife et al. 2023), we do not understand 
many aspects of photosynthetic energy conversion, such as de-
tails of its evolution, dynamics, and regulation.
In this commentary, a number of researchers working on fun-
damental aspects of photosynthesis pose and discuss what they 
view as the most compelling open questions in their areas of re-
search. We begin at the reaction center and catalytic environment 
of the Mn cluster and move outwards to address questions related 
to light harvesting, photosystem evolution, thylakoid membrane 
dynamics, and alternative electron flow pathways. We have 
avoided questions related to Rubisco, as these have been covered 
in many recent reviews and perspectives (Carmo-Silva and 
Sharwood 2023, and references cited therein; Croce et al. 2024). 
Two final sections address the link between photorespiration 
and stomatal control and adaptive changes that make an enzyme 
suitable for its function in the C4 carbon-concentrating pathway 
present in important crop species.
How does the catalytic environment of the 
Mn4CaO5 cluster accommodate and control 
the multielectron/multiproton reaction 
during water oxidation?
(Written by Vittal K. Yachandra, Jan Kern, and 
Junko Yano)
The model for photosynthetic water splitting proposed by Kok 
et al. (1970), based on the yield of dioxygen as a function of light 
flashes, has served as a blueprint for all subsequent work on the 
mechanism of the water-splitting reaction. This kinetics-based 
model proposed the advancement of the oxygen-evolving 
complex (OEC), the site where the oxidation of water occurs in 
photosystem II (PSII), through five intermediates known as the 
S-states (Si, i = 0 to 4). The absorption of four photons, one at a 
time, advances the OEC from S0 through S4, after which O2 is re-
leased. This model, called the Kok S-state cycle or clock 
(Fig. 1A), provided an elegant explanation for the coupling of the 
one-electron photochemistry occurring at the PSII reaction center 
to the four-electron redox chemistry occurring at the OEC. The 
working hypothesis was that upon the absorption of a photon, 
an oxidation equivalent is stored on the OEC, and after reaching 
the S4 state, the most oxidized intermediate, two water molecules 
are oxidized to form one molecule of O2. The S0, S1, S2, and S3 
states are stable and can be trapped, but the S4 state is normally 
referred to as a short-lived or transient state that appears during 
the S3 to S0 transition prior to O–O bond formation and release of 
O2. The Kok cycle also entails the release of four protons and four 
electrons. The consensus is that the protons are released in a 1, 0, 
1, 2 pattern for the S0 to S1, S1 to S2, S2 to S3, and S3 to [S4] to S0 
transitions, and the electrons extracted from the donor OEC site 
by the PSII reaction center are transported via the electron trans-
fer components to the quinones on the acceptor side.
Identifying the structure of PSII and the Mn-containing cata-
lytic metal cluster of these intermediates, especially, during the 
S3-[S4]-S0 transition has been challenging and the subject of 
many studies (reviewed in (Wydrzynski and Satoh 2005; Nelson 
and Yocum 2006)) using diverse spectroscopic techniques, such 
as X-ray (Yachandra et al. 1996; Dau and Haumann 2008; Yano 
and Yachandra 2014), Fourier Transform Infrared Spectroscopy 
(FTIR; Debus 2015; Nagao et al. 2018), electron paramagnetic 
resonance (EPR; Lubitz et al. 2019), and X-ray crystallography 
Figure 1. The Kok cycle and structure of PSII. A) The Kok cycle is shown in the middle, with the structures from XFEL crystallography of the S-states and 
at intermediate time-points between the S2 and S3, S3 and S0 states. O5 is shown in blue, and in red is the insertion of OX between Mn1 and Ca during S3 
to S0 and its disappearance between the S3 and S0 transitions. The sigma value in crystallography is used for the scaling of the electron density; a higher 
value shows higher accuracy. B) On top is the structure of PSII, showing the electron transfer components, the OEC, and the proton and water channels 
(in red, blue, and green) in one monomer (left). The transmembrane helices and the helices in the extrinsic polypeptides are shown in color in the 
monomer on the right. Below this is the oxo-bridged Mn4Ca cluster in the S1 state in detail, with the water and other terminal ligands of Mn and Ca 
provided by the protein glutamate, aspartate, and histidine residues.
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(Umena et al. 2011). These studies revealed that the catalytic met-
al site consists of an oxo-bridged Mn4Ca hetero-nuclear cluster 
(Fig. 1B) that accumulates charge as it proceeds through the 
S-state cycle, reinforcing the hypothesis that the S0 through S3 
transitions are primarily where the oxidizing equivalents are 
stored, with little or no specific chemistry related to the O–O for-
mation from two water molecules. However, it was shown that 
during progression from S0 through the S3 state, the oxidation 
equivalents or the charge is not centered only on the Mn but 
more delocalized onto the ligands (Glatzel et al. 2004, 2013). 
There was no evidence for the presence of a peroxo or other inter-
mediates, which are intermediate species one can expect for a 
water oxidation reaction. Intermediates such as metal-peroxo, 
-oxo, and -superoxo species have been detected in the study of in-
organic water-oxidation catalytic systems. These data indicated 
that the entire chemistry involving water and the O–O bond for-
mation occurs during the last step. This is the discharge/reduction 
step, where the most oxidized S4 state is reduced to S0 state (most 
reduced) with concomitant oxidation of water either in one fell 
swoop or in a stepwise manner, in which case it would proceed 
through the expected intermediates of water oxidation, with the 
most likely being two two-electron steps resulting in dioxygen 
via a transient peroxo intermediate.
The recent advent of X-ray free-electron laser (XFEL)-based 
room temperature crystallography and X-ray spectroscopy has 
had a profound influence on the structural studies and on the 
study of the mechanism of the water-splitting reaction (Kern 
et al. 2012, 2013, 2014; Suga et al. 2015). The structures of the S0, 
S1, S2, and S3 states have been determined at ∼2 Å resolution 
(Fig. 1B). Furthermore, it is now possible to follow the structural 
changes between the S2 and S3states (Ibrahim et al. 2020) where 
the insertion of a new water occurs between Mn1 and Ca (likely 
as –OH after releasing one proton), and most importantly between 
the S3 and S0 states where the accumulated oxidizing equivalents 
are reduced and O2 is released (Fig. 1B) (Bhowmick et al. 2023).
Several events accompany the S3 to S0 transition that resets the 
Kok-clock to the S0 state, which includes the release of two pro-
tons, the release of dioxygen, and the recovery of one water at 
the OEC. The newly introduced ligand between Mn and Ca in the 
S3 state disappears during the S3 to S0 step, and this is related to 
the release of O2. The release of two protons signaled by changes 
in the gate region in the proton channel (Hussein et al. 2021) have 
been observed at two different time-points in the XFEL crystallog-
raphy data (Bhowmick et al. 2023). Yet, it is not clear how exactly 
O–O formation and the recovery of the catalytic center takes 
place, due to the resolution of the current crystallography data 
that hinders accurate modeling of oxygen positions in the fraction 
of the changing population. The reduction process may create a 
longer lived intermediate, perhaps a peroxo species, which is 
two-electron reduced from the most oxidized S4 state. This is 
one step away from a further two-electron reduction of the OEC 
that results in the release of O2. We await structural studies at 
higher resolution.
The ability to visualize PSII structural changes through the cat-
alytic process and understand the importance of the overall pro-
tein structure in the water-oxidation mechanism has also 
opened up new questions that are perhaps foundational to 
many multielectron catalytic reactions. How does the protein en-
vironment modulate the structure to neutralize the charge den-
sity changes on the OEC during catalysis? How does the water 
network, together with the amino-acid residues, facilitate proton 
transfer and substrate transport? How are the reaction kinetics af-
fected by different environmental parameters such as pH and 
temperature? And how does the fundamental mechanism of the 
OEC remain the same through the diverse evolutionary tree 
(Hussein et al. 2023)? Recent advances in cryogenic electron 
microscopy in combination with site-specific mutations will likely 
contribute to answering some of these questions. While 
Thermosynechococcus vestitus (previously known as T. elongatus) 
has often been used for crystallography due to the availability of 
high-quality crystals, more species are being used for structural 
studies with cryogenic electron microscopy, and the structures 
from different organisms have been compared. Such comparisons 
can help to address questions about the fundamentally important 
design components for the water oxidation reaction in nature. In 
addition, the above-mentioned XFEL crystallography at room 
temperature, along with various spectroscopic methods, enables 
us to bridge structural information with spectroscopic data and 
provide the spatial and temporal sequence of events during the re-
action. Thus, the study of the light-driven water oxidation reac-
tion in PSII provides a unique opportunity to understand how 
the metal catalytic center and protein environment orchestrate 
and enable the multielectron/multiproton reaction, providing 
fundamental knowledge that goes beyond the water oxidation 
reaction.
How do protein networks control light 
harvesting?
(Written by Gabriela Schlau-Cohen and Dvir 
Harris)
In photosynthetic light harvesting, a network of chromophore- 
containing antenna proteins absorbs light and transports photo-
energy over distances of tens to hundreds of nanometers to reach 
reaction centers (Fig. 2). Despite the long distances, this transport 
can occur with near-unity quantum efficiency. At the same time, 
the efficiency is actively regulated by the network in a photopro-
tective response to light levels (Eberhard et al. 2008). The function-
ality of the network relies on the collective behavior of the 
constituent proteins. Yet studies have typically been limited to in-
dividual proteins, where relevant properties are often absent, or 
intact systems, where contributions from many proteins are com-
bined to obscure behaviors of interest (van Grondelle and 
Novoderezhkin 2006). Resolution of the interactions and dynam-
ics between proteins has been a long-standing challenge, leading 
to the major open question: How do protein networks control light har-
vesting? We discuss three facets of this open question.
How is energy transferred between proteins?
The efficient transport of photoenergy through the antenna com-
plexes to the reaction center is well established in photosyn-
thetic organisms. Such long-distance transport emerges from a 
series of energy transfer steps within photosynthetic mem-
branes. Cumulatively, these steps contribute to the hundreds 
of picoseconds timescale of solar energy conversion, whereas 
the relaxation time within individual proteins is approximately 
only one picosecond. The vast majority of previous experiments 
focused on individual proteins, even though transfer between 
proteins occupies most of the total time and is the key to long- 
distance energy transport (Fiebig et al. 2023). Despite its crucial 
importance, protein-to-protein energy transfer has been chal-
lenging to measure.
Several questions remain about how protein-to-protein energy 
transfer enables robust long-distance transport. In green plants, 
the energetic gradient of the protein network towards the reaction 
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enter is nearly flat (Mirkovic et al. 2017). How a nearly flat gradient 
produces an effective directional energy flow remains unclear. 
Both the composition and organization of the protein network 
also change under light conditions. Energy transfer is highly sen-
sitive to distance, and so even distance changes of a single nano-
meter can change the rate by an order of magnitude. Despite this 
sensitivity, the quantum efficiency of light harvesting changes 
minimally—less than 10%—with network organization (Fleming 
et al. 2012). Thus, how network reorganization impacts energy 
transport is not fully understood.
Despite the inherent challenges, several promising directions 
can now probe protein-to-protein energy transfer. Because of 
spectroscopic improvements, energy transfer rates have been 
extracted for spectrally shifted antenna proteins in membrane 
fragments or in vivo (Tiwari et al. 2018; Lüer et al. 2012). 
Alternatively, model membrane systems have been used to bio-
chemically isolate antenna proteins for measurements of 
protein-to-protein energy transfer (Dai et al. 2018; Wang et al. 
2023a, 2023b, 2023c). Finally, an integrative approach using 
single-molecule spectroscopy and cryogenic electron microscopy 
probed the heterogeneity associated with antenna to reaction 
center energy transfer (Harris et al. 2023). These recent technolog-
ical advances enable investigation of the important and funda-
mental process of energy transfer between proteins.
How does the network regulate photoprotection?
In oxygenic photosynthesis, excess energy can cause photooxida-
tive damage to proteins. Under high light conditions, a drop in the 
lumenal pH activates a series of photoprotective processes, 
known as nonphotochemical quenching (NPQ). The fastest NPQ 
process is qE, the dissipation of excess energy as heat. The major 
antenna protein, light-harvesting complex II (LHCII), is a major 
site of dissipation for PSII (Schlau-Cohen et al. 2015; Nicol and 
Croce 2021), and its organization within the protein network 
changes, notably through the formation of clusters (Goral et al. 
2012; Ruban and Johnson 2015). Recapitulating the physiological 
LHCII–LHCII interactions responsible for the functional switch 
has remained challenging, and so how these interactions regulate 
the dissipative pathways of photoprotection in vivo remains 
unclear.
Studies on aggregates and arrays of LHCII, which aim to emulate 
LHCII clustering, have shown enhanced dissipation, and a pH de-
pendence reminiscent of the in vivo conditions (Barros and 
Kühlbrandt 2009; Petrou et al. 2014; Son et al. 2021). A dissipative 
photophysical pathway, chlorophyll-to-carotenoid energy transfer, 
was recently clearly resolved with the cluster and pH dependence 
expected for photoprotection. In recent in vivo measurements, 
the carotenoid photophysics correlated with NPQ (Park et al. 
2019), suggesting that the photoprotective switch involves these 
important biomolecules. Such effects may be induced by LHCII 
clustering.
What are the protein–protein interactions that 
govern network architecture?
In recent years, the organization of the protein networks has been 
revealed in exquisite detail (Wietrzynski et al. 2020; Opatíková 
et al. 2023; Feng et al. 2023). Advances in structural biology, 
including electron microscopy and tomography at cryogenic tem-
peratures and atomic force microscopy, have provided snapshots 
that dramatically advanced our understanding. However, the in-
teractions between the proteins that drive the formation of these 
networks and their reorganization in response to environmental 
stimuli remain unknown. While a small number of computational 
studies have investigated these interaction energies (Schneider 
and Geissler 2013; Mao et al. 2023), the well-established deviation 
between computational energies and experimental benchmarks 
(Keskin et al. 2016) means that new methods are needed to deter-
mine the thermodynamic driving forces behind protein networks. 
Recent experimental measurements of the interaction energies 
between LHCII revealed a net attraction of ∼−5 kBT, which repre-
sents an enthalpically dominated thermodynamic driving force 
behind LHCII clustering. Such studies capture the interactions 
Figure 2. PSII from green plants. Chromophore-containing antennae capture and transport light energy (orange) to the PSII reaction center, where 
charge separation occurs (black). Under high light conditions, the local cellular environment and the organization of the protein network activate 
photoprotection within the antenna proteins, likely involving a switch of the photophysics of the embedded chlorophyll and carotenoids associated 
with different structural states (PDB: 5XNM). The protein network is formed through van der Waals interactions between proteins.
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behind the membrane organization of PSII from the perspective of 
equilibrium statistical thermodynamics, which has a long and 
rich tradition in biology (Manna et al. 2023).
In conclusion, structural and spectroscopic advances over the 
past decades built an understanding of the function and composi-
tion of the protein networks of photosynthetic light harvesting. In 
particular, the structure, organization, and dynamics of the con-
stituent proteins have been revealed. The open questions sit at 
the interface between these proteins. Resolution of the interac-
tions between proteins and how these interactions modulate 
function requires further improvements to our tools that charac-
terize and manipulate these biosystems. Answering the central 
question—How do protein networks control light harvesting?— 
remains a grand challenge for the field.
Why do diatoms have so many different 
light-harvesting proteins?
(Written by Nicolas Herrmann and Claudia 
Büchel)
Vascular plants express an array of light-harvesting complexes 
(LHC) and the different functions of these LHCs as major or minor 
antennae for PSII, as PSI-specific antennae, or as mobile antennae 
in the context of state transitions are well described (Croce and 
van Amerongen 2020). Algae, by contrast, contain many more Lhc 
proteins (Lhc and LHC typically are used to denote the algal and vas-
cular plant proteins, respectively), and one of the intriguing ques-
tions is the reason behind this diversity. Besides the well-studied 
green algae, there are many algal groups that use different pigments 
and even different proteins for light harvesting (Büchel 2015). Most 
prominent are the Stramenopiles, which are distantly related to 
green algae and vascular plants. These algae are derived from a so- 
called secondary endosymbiosis, where a eukaryotic host engulfed 
an organism related to red algae that was evolutionarily reduced to 
the chloroplast (Bhattacharya and Medlin 1995). The group of 
Stramenopiles with the best-studied light-harvesting systems are 
diatoms, unicellular organisms that are responsible for 20% of pri-
mary production worldwide (Field et al. 1998). Diatoms are found 
in many different habitats, ranging from biofilms to free-living 
marine or limnic planktonic forms. Marine diatoms are relatively 
easy to culture, and after whole-genome sequences as well as 
molecular tools for transformation became available, different 
species emerged as reference systems, i.e. the pennate diatom 
Phaeodactylum tricornutum and the centric diatoms Thalassiosira pseu-
donana, Cyclotella meneghiniana, and Chaetoceros gracilis.
Diatoms possess a photosynthetic apparatus that is similar to 
that of vascular plants, including membrane-intrinsic antenna 
proteins of the Lhc family (Büchel 2015). However, their pigmenta-
tion is very different. Whereas chlorophyll (Chl) a is present in di-
atoms, Chl b is missing and only 2 to 3 molecules of Chl c are bound 
instead. The carotenoid fucoxanthin (Fx) is the main accessory 
pigment with up to seven molecules per protein of the eponymous 
Fucoxanthin-Chlorophyll Proteins (FCP). The Fx molecules are 
bound to various sites of FCP, inducing differences in absorption 
(Fig. 3A). Thus, FCP absorbs up to 585 nm, giving rise to the char-
acteristic brown color of diatoms and optimizing photosynthesis 
in deeper water, where blue-green light dominates.
A wealth of spectroscopic data exists about the excitation en-
ergy transfer inside FCP complexes. Fx displays efficient excitation 
energy transfer to Chl a (Papagiannakis et al. 2005). The fastest 
transfer (∼30 to 60 fs) takes place between Chl c and Chl a 
(Butkus et al. 2015). Thus, excitation energy transfer between dif-
ferent Chl molecules is faster and FCP rely more on carotenoid ab-
sorption for light-harvesting than do Lhc of vascular plants. 
Nonetheless, these features are similar for all Lhc proteins from 
diatoms studied so far and do not shed light on a possible func-
tional heterogeneity.
All diatom species contain a much higher number of expressed 
Lhc genes than found in vascular plants. T. pseudonana has more 
than 30 FCP proteins, C. meneghiniana has 23 FCPs, and C. gracilis 
and P. tricornutum have 22 and 32 FCPs, respectively (Kumazawa 
et al. 2022). PSI of diatoms is surrounded by monomeric FCPs, 
but the antenna is much larger than that of vascular plants, com-
prising ∼16 to 24 different FCPs in the case of C. gracilis (Nagao 
et al. 2020; Xu et al. 2020) (Fig. 3B). PSII structures are available 
from three centric species that differ tremendously in their FCP 
arrangement and composition, a diversity completely unknown 
in vascular plants. Whereas C. gracilis has three monomeric and 
two tetrameric FCP per core monomer (Pi et al. 2019), PSII of 
C. meneghiniana (Wang et al. 2023a, 2023b, 2023c) has only six 
FCP proteins. The high-resolution structure of T. pseudonana PSII 
shows one additional FCP (Feng et al. 2023) and is in contrast to 
the low-resolution structures, where three monomers and one 
trimer were modeled (Arshad et al. 2021). In addition, even the oli-
gomeric state of FCP complexes found in the free pool of FCPs dif-
fers between species: for P. tricornutum Lhcf4 homodimers (Wang 
et al. 2019) and different trimers (consisting of the other FCP 
Figure 3. The FCP complex. A) Absorption spectrum of an FCP complex (black, FCPa from C. meneghiniana) and its pigments. Fx molecules are bound at 
different sites, absorbing more to the red wavelength range (Fx “red”), intermediate (Fx “green”), or to the blue (Fx “blue”). B) Scheme of FCP 
arrangement in PSI, PSII, and the free FCP pool of different species. Lhcx proteins are shown in red, other FCPs are colored according to their subfamily 
(light purple, Lhcf; blue, Lhcr; purple, lhcq; gray, unknown). For references, see text.
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proteins including Lhcf4) (Gundermann et al. 2013) were reported, 
whereas analysis of C. meneghiniana revealed trimers and 
nonamers (Röding et al. 2018; Wang et al. 2023a, 2023b, 2023c). 
Oligomerization influences the pigment network and correspond-
ingly, excitation energy transfer into the reaction centers. So far, 
no functional reason is known for this huge inter-species 
diversity.
To cope with rapid changes in light intensity while maintaining 
optimum photosynthetic energy flux, diatoms employ nonphoto-
chemical quenching (NPQ), which is very fast and has a high ca-
pacity compared to NPQ of vascular plants (Ruban et al. 2004). 
NPQ is triggered by a high trans-thylakoid pH gradient under 
strong light and relies on a xanthophyll cycle that converts diadi-
noxanthin (Ddx) to diatoxanthin (Dtx) under excess light (Lavaud 
et al. 2002) and special FCP proteins called Lhcx. Diatoms contain 
many Lhcx proteins. Lhcx1 was demonstrated to be involved in 
NPQ in P. tricornutum (Bailleul et al. 2010) as well as in C. meneghini-
ana (Ghazaryan et al. 2016), but Lhcx1 cannot sense pH (Buck et al. 
2021) and therefore the mechanism triggering NPQ remains 
enigmatic. Aggregation of FCPs containing Lhcx1 from the C. mene-
ghiniana FCP pool reduced fluorescence in vitro and Dtx enhanced 
this fluorescence, which resembles NPQ in vivo (Gundermann and 
Büchel 2012). Domain swap experiments demonstrated the im-
portance of a Trp residue of Lhcx1 that is close to the presumed 
Ddx binding site (Buck et al. 2021), hinting at a direct connection 
between Lhcx proteins and Ddx/Dtx. However, although the par-
ticipation of Dtx in NPQ is well established, an in-depth spectro-
scopic analysis will be required to prove that Dtx that is bound 
to Lhcx1 participates in NPQ. The centric diatoms T. pseudonana 
and C. meneghiniana contain a special Lhcx protein, Lhcx6_1, 
which was found to be located in PSII complexes, harbors Dtx, 
and might be the second quenching site usually found in centric 
diatoms (Calvaruso et al. 2020, Grouneva et al. 2008). The sole 
presence of Dtx in an Lhcx structure does not prove its involve-
ment in NPQ. Various abiotic stresses were shown to regulate 
the expression of the other three Lhcx proteins in P. tricornutum 
(Taddei et al. 2016), tuning NPQ. This functional diversification 
might help diatoms cope with variable environments.
Our understanding of the arrangement of FCPs around the pho-
tosystems, pigment organization, and excitation energy transfer 
has greatly improved in recent years. But why is FCP organization 
around PSII so diverse among the different species and what are 
the roles of the different FCP proteins? Involvement of FCPs in reg-
ulatory functions is the obvious idea to explain the huge diversity 
in and between species, but whether this explanation is true re-
mains mostly enigmatic, as plasticity in the spatial arrangement 
of different FCPs remains unreported. And how precisely are the 
different Lhcx proteins involved in NPQ? Answering these ques-
tions will help us to understand the huge ecological and evolu-
tionary success of diatoms.
New perspectives on the evolution of the 
photosystems: Are the photosystems 
evolvable?
(Written by Tom Dongmin Kim, Emma Chaloner, 
and Tanai Cardona)
How and when photosynthesis originated remain compelling 
questions in the Life and Earth Sciences. How photosystems, the 
multicofactor and multiprotein assemblies that power primary 
production, were put together from their constitutive parts and 
for what original function, are questions that still need answers. 
The photosystems are well over three billion years old and are po-
tentially amongst the oldest enzymes (Oliver et al. 2023). On the 
one hand, that such endurance is possible suggests that photosys-
tems have an in-built capacity for evolution and adaptability that 
enable photochemical processes to occur in as varied and dynam-
ic conditions as can be found on Earth’s photic zones, and on a 
planet that has gone through remarkable transformations over 
the eons. On the other hand, photosystems are highly conserved 
and slowly evolving enzymes, often perceived as somewhat im-
mutable, even dubbed “frozen metabolic accidents” (Shi et al. 
2005). Given that all photosystems share a common origin, evi-
dence for the photosystems” evolvability is most conspicuously 
noted when comparing PSII and its unparalleled capacity to split 
water by generating some of the most oxidizing species in biology, 
with PSI, which has evolved to generate some of the most reducing 
ones.
These evolutionary considerations trigger a question that has 
received little explicit attention in the field of photosynthesis re-
search: are the photosystems evolvable? Given that photosystems 
are biological systems, it is safe to assume that they are. 
However, the question raises a number of additional fundamental 
considerations about how photosystems have changed through 
time. For example, what are the molecular mechanisms that 
make the photosystems adaptable? How quickly can a photosys-
tem gain a new function given adequate selective pressure? 
And, given selective pressures not found in nature, what new pho-
tochemical and catalytic properties could a photosystem evolve? 
We could go one step further: Can photosystems be evolved in the lab? 
We propose here that in an endeavor to answer these questions, 
we could gain new insights into why the photosystems evolved 
to be the way they are and open new pathways to greener chem-
ical and biotechnological processes.
Photosystems are modular and this modularity is the basis for 
their capacity to remain adaptable. In the wild, this means that 
the photochemical or catalytic properties of a photosystem can 
be modified to respond to the environment by exchanging subu-
nits, which work as replaceable modules. The better-known ex-
ample of this is the fine-tuning of PSII energetics to perform 
optimally under different light intensities, and the most extreme 
example is the transformation of PSII from a water-oxidizing to 
a Chl f-producing enzyme (reviewed recently by Oliver et al. 
(2023)). In both cases, these functional alterations are achieved 
by replacing a standard D1 subunit for a variant form of D1. And 
in both cases, PSII evolved new photochemical or catalytic proper-
ties by acquiring modifications on a spare copy of a gene encoding 
the D1 subunit within the genome of cyanobacteria. Alterations of 
PSI by a similar mechanism are also known. For example, some 
cyanophages carry a set of PSI genes that are deployed upon infec-
tion of the cyanobacterium. In this adaptation, the standard PsaF 
and PsaJ subunits are replaced by a subunit encoded in the cyano-
phage’s genome which is a fusion of the two (Sharon et al. 2009). 
This novel subunit has features that make the complex more pro-
miscuous to protein electron carriers (Mazor et al. 2014). Another 
adaptation commonly observed in heterocystous cyanobacteria 
and their close relatives is the exchange of PsaB paralogs, which 
is likely to lead to a fine-tuning of the energetics of the complex 
in ways and conditions that are yet to be understood (Gisriel 
et al. 2023). Some strains encode up to four distinct PsaB versions. 
While the exchange of just a single subunit is sufficient to modify 
the photochemical and catalytic properties of the photosystems, 
there is no appreciable boundary as to the extent of change that 
can occur driven by sustained evolutionary pressures. A clear 
example of this is the far-red light photoacclimation response 
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(Gan et al. 2014), which involves the swap of at least five subunits 
of PSII, and six of PSI, and the use of three distinct chlorophyll 
types to enable oxygenic photosynthesis under far-red light and 
in the absence of visible light.
The next step towards understanding the evolution of the pho-
tosystems is to evolve them in a laboratory setting and attempt to 
change their catalytic and photochemical properties purpose-
fully. There is experimental evidence suggesting that the modu-
larity of PSII can facilitate the application of directed evolution 
methods. For example, random mutagenesis of D1 was used to se-
lect for PSII complexes more tolerant to high light intensities 
(Narusaka et al. 1999), with tolerance to herbicides (Narusaka 
et al. 1998), and with tolerance to ionizing radiation (Rea et al. 
2011). However, these studies stopped at a single round of muta-
genesis and selection. In addition, directed evolution was used 
to successfully change the directionality of electron transfer of 
the purple bacteria reaction center (Faries et al. 2012).
Evolving PSII could involve modifications of the electron donor 
side to change the nature of the water-oxidizing cluster and to en-
able access to other substrates (Fig. 4A). For example, to study one 
of the plausible transitional stages in the origin of PSII (Chernev 
et al. 2020), it may be possible to evolve a PSII that can efficiently 
oxidize Mn but cannot split water. Alternatively, a PSII could be 
evolved to decompose a complex organic pollutant in a sequential 
multielectron multiproton extraction process. The energetics of 
charge separation could be optimized for the specific catalytic de-
mands imposed by the new substrates. There is evidence support-
ing the idea that PD1, one of the key redox chlorophylls of PSII that 
has a midpoint potential of about +1.2 V, is naturally tuned down 
by −0.14 V to compensate for electrostatic effects created by the 
Mn4CaO5 cluster (Ishikita et al. 2006), implying that the photo-
chemical core could be pushed to generate even higher oxidizing 
potentials if desired. Evolving PSI could involve adding novel cat-
alytic domains, which could be further optimized with directed 
evolution methods. In this sense, PSI could be engineered to trans-
fer electrons from the terminal iron–sulfur clusters to the cofac-
tors of the new domains, resulting in novel photobio catalysis 
(Fig. 4B). This concept has been demonstrated in the genetic fu-
sion of the extrinsic subunits of PSI with hydrogenase enzymes 
(Appel et al. 2020; Kanygin et al. 2020; Wang et al. 2023a, 2023b, 
2023c). The chimeras were all characterized as functional photo-
systems capable of light-driven H2 evolution. Such experiments, 
while opening new avenues for applications, would also test 
many existing rationales aimed at explaining photosystem struc-
ture and function. For example, what are the exact energetic 
tradeoffs between photochemical efficiency and photoprotection? 
Thus, if oxygen were not to be a reaction by-product of a hypothet-
ical evolved PSII oxidizing a new substrate, could energy that is 
sacrificed by the reaction center to prevent back-reactions be re-
directed towards improving turnover efficiency? If that were the 
case, then, what would be the maximum rate of water oxidation 
of PSII, i.e. if the system had not needed to evolve into its bioener-
getics protective mechanisms against the production of reactive 
oxygen species? By exploring these questions, a positive feedback 
loop can be created between research aimed at understanding 
photosystem evolution and developing new photosystem-based 
technologies for better biocatalysis.
To conclude, two specific questions can be asked to guide new 
research. First, what are the limits of photosystem evolvability? 
The answer to this question will help us understand the limits 
of light-dependent life on Earth and extrasolar planets in habit-
able zones. It can also help us understand why life came to be 
the way it is. Second, how can novel light-driven enzymes, featur-
ing customized photochemical and catalytic properties, contrib-
ute to making the chemical and biotechnological industries 
more sustainable? This question is an invitation to a new genera-
tion of photosynthesis researchers to use their creativity in ex-
ploring how new photosystems could be harnessed as solutions 
to some of the global challenges we face.
Thylakoid membrane dynamics: How are 
protein biogenesis and mobility 
orchestrated in this densely packed system?
(Written by Conrad Mullineaux)
Thylakoid membranes in chloroplasts and cyanobacteria are 
densely packed with protein complexes. They present a crowded 
environment, not only in the plane of the membrane but also in 
the third dimension, where parallel membrane surfaces may be 
Figure 4. Conceptualization of evolved photosystems. A) A PSII evolved to bind a dinuclear Ru cluster (Ru2O2), which catalyzes a hypothetical 
specialized oxidative reaction. B) A hypothetical chimeric PSI fused with a reductive dehalogenase and evolved for optimal electron transfer to the 
catalytic site at a cobalamin cofactor (B12). Protein scaffolds of the photosystems are shown in transparent gray ribbons, and that of the reductive 
dehalogenase in magenta; chlorophylls are shown in green, carotenoids in purple, and iron–sulfur clusters are depicted with yellow and orange 
spheres. PSII structure was modified from PDB ID 3WU2, PSI from 1JB0, and reductive dehalogenase from 5M2G.
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tightly appressed, or in close proximity and sandwiching a dense 
mass of soluble protein. Dense packing of the system allows a 
high concentration of pigment-protein complexes, which is neces-
sary for the efficient absorption of sunlight. However, it creates 
challenges for other aspects of membrane function, including reg-
ulatory re-organization, diffusion of mobile electron carriers, and 
the biogenesis of protein complexes, which could all be impeded 
by the crowded environment in and around the thylakoids 
(Kirchhoff 2014a).
The translation and membrane insertion of membrane- 
integral protein complexes invariably seem to occur in specialized 
regions of the membrane that may be quite remote from much of 
the mature functioning thylakoid surface. This likely arises be-
cause the sheer bulk of the ribosomes excludes them from most 
of the crowded thylakoid system. In cyanobacteria, ribosomes 
are found only at the proximal thylakoid surface adjacent to the 
central cytoplasm and are therefore excluded from about 90% of 
the thylakoid system (Fig. 5; Rast et al. 2019; Mahbub and 
Mullineaux 2023). mRNAs encoding membrane-integral thylakoid 
proteins are found at the same membrane surface (Mahbub et al. 
2020; Mahbub and Mullineaux 2023). In chloroplasts of the green 
alga Chlamydomonas, translation is heavily focused on a special-
ized “T-zone” surrounding the pyrenoid (Schottkowski et al. 2012). 
In vascular plant chloroplasts, ribosomes are excluded from the 
tightly appressed grana membrane surfaces, but ribosomes also 
appear very scarce across large areas of the stromal lamellae, 
being concentrated at the most peripheral regions of the thylakoid 
that have unimpeded access to the chloroplast stroma (Shimoni 
et al. 2005).
If translation at the thylakoid is highly localized, it raises the 
question of how newly synthesized protein complexes can find 
their place in the mature thylakoid membrane. Thylakoid mem-
branes seem to be more interconnected than is often apparent 
at first sight. For example, the thylakoids of the cyanobacterium 
Synechococcus elongatus approximate a set of nested cylinders 
(Fig. 5), but electron tomography revealed membrane bridges 
that interconnect the whole system (Nevo et al. 2007). So, it is 
likely that there is a continuous membrane surface connecting 
every region of a thylakoid to a translation zone, but the problem 
of restricted protein diffusion in the crowded thylakoid mem-
brane environment remains—is it feasible for a newly synthesized 
complex to diffuse from the site of its translation to the furthest 
parts of the thylakoid system? Taking S. elongatus as an example, 
Fluorescence Recovery after Photobleaching (FRAP) measure-
ments on GFP-tagged photosynthetic complexes suggest long- 
range lateral diffusion coefficients of the order of 10−10 cm2s−1 
(Casella et al. 2017). Such a diffusion coefficient would permit a 
complex to diffuse on average 1 µm from its starting point in 
25 s, however for radial diffusion from the proximal to the distal 
layers of the thylakoid system the problem is hugely compounded 
by the need for the complex to encounter one of the very occasion-
al bridges that connect the concentric thylakoid membrane layers 
(Nevo et al. 2007). There is an analogous problem in chloroplast 
appressed grana membranes, where FRAP measurements suggest 
a limited mobile pool of chlorophyll–protein complexes with a dif-
fusion coefficient of about 5 × 10−11 cm2s−1 (Kirchhoff et al. 2008), 
but there are restricted connections between the appressed 
membranes and the stromal lamellae (Mustárdy et al. 2008). 
Computational modeling combined with high-resolution data on 
3D thylakoid ultrastructure will be needed to define the extent 
of the problem.
If, as seems likely, movement of newly synthesized complexes 
to the furthest parts of the thylakoid system is slow, then what are 
the solutions and what are the functional implications? Taking S. 
elongatus (Fig. 5) as an example, two extreme scenarios could be 
envisaged for the incorporation of new complexes into the ex-
panding thylakoid membrane as the cell grows. In the first scenar-
io, new complexes stay close to their site of translation. As the cell 
grows and more complexes are synthesized, the parts of the prox-
imal membrane surface mature into fully functional thylakoid. 
New areas of biogenic membrane surface are generated to replace 
them, so that new thylakoid is created at the proximal side of the 
thylakoid system and older areas of thylakoid are gradually 
pushed towards the cell periphery as the cell grows. One piece 
of evidence against this idea is that in S. elongatus cells that are re-
generating their thylakoid system after growth in high light, new 
thylakoid membrane sacs seem to appear at the distal edge of 
the system between the existing thylakoids and the plasma mem-
brane (Huokko et al. 2021). These new membranes must presum-
ably be populated by protein complexes migrating from the 
proximal side of the system. The second scenario assumes that 
the proximal thylakoid membrane surface remains as a biogenic 
surface throughout, and complexes migrate out of it as they are 
assembled. It seems that this idea must apply at least to the re-
placement of the D1 subunit of PSII, which is rapidly turned 
over in high light (Komenda et al. 2012). Newly translated D1 pro-
teins must surely be needed for repair of damaged PSII complexes 
throughout the thylakoid system. Interestingly, there are indica-
tions that the mobility of chlorophyll–protein complexes in-
creases following high light exposure, in both chloroplasts 
(Goral et al. 2010) and cyanobacteria (Sarcina et al. 2006).
Both scenarios outlined above would predict gradients in thyla-
koid membrane protein composition depending on their distance 
from the nearest translation site. Repair and regulatory adjust-
ments to the population of photosystems (e.g. (Dietzel et al. 
2008; MacGregor-Chatwin et al. 2022) might take effect at differ-
ent speeds in different regions of the membrane, with strong func-
tional consequences. Understanding the consequences will 
require progress in cell biological approaches to thylakoid biogen-
esis, to complement the remarkable advances from biochemical 
and structural approaches (e.g. (Zabret et al. 2021)). Electron to-
mography has provided new insight into the 3D architecture of 
Figure 5. The photosynthetic protein biogenesis problem, illustrated by 
a schematic cross-section of a cell of the cyanobacterium Synechococcus 
elongatus. Translation of membrane-integral photosynthetic proteins 
occurs only at the proximal thylakoid surface facing the central 
cytoplasm and the nucleoid. How do new complexes get to the more 
distant parts of the thylakoid system?
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thylakoids, as in (Shimoni et al. 2005; Nevo et al. 2007; Engel et al. 
2015; Rast et al. 2019), and this should be coupled with fluores-
cence microscopy to probe membrane dynamics. Techniques 
such as structured illumination microscopy, as in (Wood et al. 
2018), will allow higher resolution than was previously achievable. 
It would be exciting to develop fluorescent labeling methods that 
distinguish between newer and older photosystems. This could 
reveal the flux of the membrane system over time, as the cell or 
organelle expands and divides. How do the dynamics of the mem-
brane and the locations of membrane protein production con-
strain photosynthetic function, and how much do they restrict 
acclimation to new conditions? These will be overarching re-
search questions for the next years.
How fluid are thylakoid membranes?
(Written by Wojciech Wietrzynski and Benjamin 
D. Engel)
One of the most intriguing characteristics of thylakoid mem-
branes is their diverse molecular architecture (Perez-Boerema et 
al. 2024). In cyanobacteria, as well as the chloroplasts of red algae 
and glaucophytes, thylakoids are decorated with bulky phycobili-
some light-harvesting antennae, which space the membranes 
apart and prevent stacking interactions. Here, PSII has been ob-
served to form linear arrays, bound underneath rows of phycobi-
lisomes (Rast et al. 2019; Li et al. 2021a, 2021b). PSI has been 
observed to bind alongside these PSII arrays in red algae (You 
et al. 2023), or in cyanobacteria to form trimers or tetramers occu-
pying separate domains within the nonstacked thylakoids 
(MacGregor-Chatwin et al. 2017).
In contrast, thylakoid architecture in the chloroplasts of green 
algae and plants can generally be subdivided into appressed and 
nonappressed (stacked and unstacked) domains (Mustárdy et al. 
2008). These chloroplasts lack phycobilisomes and instead rely 
on light-harvesting complexes (LHCs) and related membrane- 
integral antenna proteins, which do not space the thylakoid 
membranes apart. In fact, there is evidence that interactions 
between LHCII proteins contribute to membrane stacking in 
plants (Standfuss et al. 2005; Daum et al. 2010; Zeno et al. 2022). 
Stacking enforces lateral heterogeneity in thylakoid membranes: 
PSI and ATP synthase populate the nonappressed domains, 
whereas PSII is restricted to the appressed domains, and only cy-
tochrome b6f is abundant in both domains (Andersson and 
Anderson 1980; Wietrzynski et al. 2020). This segregation of com-
ponents prevents energy spillover from PSII to PSI, and has deep 
implications for the regulation of photosynthesis. Balancing of 
PSI and PSII excitation via migration of LHCII between domains 
(state transitions; Allen and Forsberg 2001), repair of PSII com-
plexes damaged by photoinhibition (Barber and Andersson 
1992), and integration of newly synthesized proteins into the 
appressed membranes would all require the exchange of 
protein complexes between the stacked and unstacked thylakoid 
regions. Therefore, to respond to varying environmental condi-
tions that chloroplasts experience throughout a typical day, thy-
lakoids should undergo some structural changes, at least at the 
scale of single protein complexes moving between membrane 
domains.
But how fluid are these thylakoid membrane domains? In other 
words, how mobile are the protein complexes and lipids within 
the plane of the membrane under short timescales? Do the com-
ponents freely mix like a liquid, or are they more static like a gel? 
While plastoquinone must diffuse between PSII and cytochrome 
b6f, it is unknown whether the larger photosynthetic complexes 
freely move around within their resident membrane domain or 
are fixed in space and experience only small oscillations 
(Kirchhoff 2014a). On the one hand, the low saturation of thyla-
koid lipids should facilitate the diffusion of protein complexes in 
the membrane; on the other, mobility might be restricted by mo-
lecular crowding from the high protein concentration (thylakoids 
are ∼70% protein [Kirchhoff et al. 2002]) as well as electrostatic in-
teractions between stacked thylakoids (Standfuss et al. 2005). Can 
current technology measure the degree of mobility for thylakoid 
protein complexes in vivo? And would changes in this mobility im-
pact cellular physiology and photosynthesis? Does it matter 
whether thylakoid membranes are more fluid or more static?
Two electron microscopy (EM) techniques can directly visualize 
individual protein complexes within thylakoid membranes in 
vivo. Freeze-fracture EM has been used for decades to quantify 
the organization of photosynthetic complexes within fractured 
membrane planes (Ojakian and Satir 1974; Charuvi et al. 2016). 
More recently, cryo-electron tomography has extended this visu-
alization to both sides of thylakoid membranes within the cell, 
providing a clear view of the lateral heterogeneity between mem-
brane domains (Wietrzynski et al. 2020). However, these EM ap-
proaches can only provide frozen snapshots of protein complex 
organization, begging the question of how mobile these com-
plexes are. Fluorescence recovery after photobleaching (FRAP) 
can provide information on protein mobility, and it has been 
used to determine that PSII complexes do not rapidly diffuse be-
tween separate grana stacks (Kirchhoff et al. 2008), which is con-
sistent with lateral heterogeneity. However, the resolution of this 
technique is far too limited to follow the mobility an individual 
PSII complexes within an appressed grana membrane. 
Spectroscopy can detect in vivo changes in the general ordering 
of thylakoid proteins, but it provides no spatial information on 
the movement of complexes. Only high-speed atomic force micro-
scopy (AFM) combines the spatial resolution to map single com-
plexes in a membrane with the temporal resolution to track 
their mobility (Clausen et al. 2014; Onoa et al. 2020). However, 
AFM is limited to measuring the topology of isolated membranes, 
which may not reproduce the thylakoid organization and fluidity 
found inside a living cell. Unfortunately, no single technique in 
use today can track the short-timescale mobility of individual pro-
tein complexes within native thylakoids.
Observations of PSII organization provide hints that this photo-
synthetic complex may experience changes in its mobility, and 
also underline the potential differences between isolated and na-
tive membranes. In cyanobacteria and red algae, PSII complexes 
bound along linear arrays of phycobilisomes must experience 
some restriction in their degrees of freedom. In isolated thylakoids 
of plants and diatoms, PSII has been observed in both a dispersed 
organization and arranged in two-dimensional pseudocrystalline 
lattices (Fig. 6) (Staehelin 1976; Dekker and Boekema 2005; 
Kirchhoff et al. 2007; Daum et al. 2010; Sznee et al. 2011; Levitan 
et al. 2019). These PSII lattices would further restrict the degrees 
of freedom, and they also interact with lattices in neighboring 
membranes to induce stacking of isolated thylakoids (Daum 
et al. 2010), limiting the mobility of PSII in two appressed mem-
branes. In vivo, only the dispersed organization of PSII has been 
observed to date, with the exception of PSII lattices within dehy-
drated resurrection plants (Craterostigma pumilum) (Charuvi et al. 
2015).
What would happen to PSII-related processes when the com-
plexes are organized in a lattice within appressed thylakoids? 
How would the lattice affect plastoquinone diffusion to and 
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from PSII? Are PSII able to leave the lattice when damaged? Are in-
itiators (e.g. kinases; Depège et al. 2003) and effectors (e.g. pro-
teases; Lindahl et al. 2000) located in the stroma able to reach 
them? Do PSII lattices receive less excitation from LHCII? If PSII 
lattices are physiologically relevant, perhaps a transition between 
mobile dispersed PSII and the static lattice configuration can help 
chloroplasts resist particularly harsh environmental conditions, 
such as dehydration of resurrection plants. The thylakoids may 
be locked in a nonphotosynthetic state that can activate again 
upon rehydration. Could such a change in thylakoid fluidity occur 
locally as a photoprotective mechanism to regulate excess excita-
tion or to sequester subpopulations of damaged PSII? Due to the 
technical challenge of observing PSII dynamics in vivo, exploring 
these questions will require approaches that integrate structural 
observations, biophysical measurements, physiology, and molec-
ular simulations.
How does alternative electron flow 
contribute to the maintenance of robust 
photosynthesis?
(Written by Arthur R. Grossman)
Light absorption drives photosynthetic electron transport and the 
synthesis of reductant and ATP to energize CO2 fixation, growth, 
and other anabolic processes. However, photosynthetic antennae 
often absorb excess excitation energy, especially upon exposure 
of plants to high/fluctuating light intensities or stress (Saroussi 
et al. 2017). To cope with excess absorbed light energy, photosyn-
thetic organisms have evolved mechanisms to dissipate this en-
ergy as heat through nonphotochemical quenching (NPQ), 
including energy-dependent quenching (qE), state transitions 
(qT), photoinhibition (qI), and zeaxanthin-dependent quenching 
that does not require a high ΔpH across the thylakoid membranes 
(qZ) (Niyogi and Truong 2013).
Photosynthetic electron transport involves linear electron flow, 
which generates ATP and reducing power, and alternative electron 
flows (AEF), including cyclic electron flow and H2O-to-H2O cycles in 
which O2 is reduced to H2O (Fig. 7). Linear electron flow does not 
generate enough ATP to sustain photosynthesis, a deficit made 
up by the various AEF pathways. Additionally, AEF deposits protons 
in the thylakoid lumen that can help establish NPQ, impact the rate 
of linear electron flow, protect PSI from damage under high/ 
fluctuating light (see the section below by Burlacot), and help 
balance the ATP:NADPH ratio, which can impact PSI donor and ac-
ceptor side regulation (Munekage et al. 2004; Allahverdiyeva et al. 
2015a, 2015b). AEF reactions include (i) Mehler-type, (ii) NADPH:fla-
vin oxidoreductase (FLV) catalyzed O2 reduction, designated pseu-
docyclic electron flow, (iii) plastoquinol terminal oxidoreductase 
(PTOX) activity, and (iv) chloroplast-to-mitochondria electron 
flow, which shuttles electrons between the organelles. The bio-
chemical shuttles important for the flow of reductant between 
the chloroplast and mitochondria translocate oxaloacetate/malate 
(OMT; Fig. 7, #1a), malate/aspartate (Dao et al. 2022), and triose-P/Pi 
(TPT) (Fig. 7, #1b) (Huang et al. 2023), coupled with redox exchange 
reactions catalyzed by malate dehydrogenase (MDH) and glyceral-
dehyde 3-P dehydrogenase (GAPDH). We describe basic features of 
these pathways, with a focus on Chlamydomonas, and highlight 
some major unanswered questions about their role(s) in maintain-
ing robust photosynthesis.
Cyclic electron flow
Cyclic electron flow (Fig. 7, #2a, 2b) directs reducing equivalents 
from the PSI acceptor to the donor side, enabling proton deposition 
in the thylakoid lumen and the use of the proton gradient for ATP 
synthesis without generating reducing equivalents. Most photo-
synthetic organisms use two cyclic electron flow pathways, one in-
volving the proton gradient generation 5 (PGR5) and the PGR-Like 1 
(PGRL1) proteins (DalCorso et al. 2008), and the other the plastid 
Type I NDH (NADPH:flavin oxidoreductase) complex in vascular 
plants (Joet et al. 2001) or Type II NDH, designated NDA2, in green 
algae (Desplats et al. 2009). The Type I NDH has 11 chloroplast- 
encoded and >19 nucleus-encoded subunits and is similar to the 
bacterial/mitochondrial NADH:UQ respiratory complex (complex 
1) (Peltier et al. 2016); it likely uses reduced ferredoxin (FDX) as 
the electron donor (Yamamoto et al. 2011). The PGR5/PGRL1 path-
way also appears to involve PetO, FNR, and ANR2 (Buchert et al. 
2018) that may function in a cyclic electron flow supercomplex 
(Joliot et al. 2022). Tobacco (Nicotiana tabacum) and Arabidopsis 
(Arabidopsis thaliana) with lesions in genes encoding chloroplast 
NDH subunits do not exhibit strong phenotypes under mild condi-
tions (Shikanai et al. 1998). The Chlamydomonas Type II NDA2 
transfers electrons from NAD(P)H to the PQ pool and becomes 
highly active during N deprivation (Saroussi et al. 2016).
Mehler reaction
The Mehler reaction (Fig. 7, #3) involves noncatalyzed O2 reduc-
tion using PSI-derived electrons (Asada 1999). It can generate reac-
tive oxygen species (ROS), including superoxides (O2
−) and 
hydrogen peroxide (H2O2) that can cause damage to various mol-
ecules of the cell, or the O2
− can be converted to H2O by the 
Dispersed (and mobile?) Organized lattice
Order &
local concentration
Mobility?
PSII ?
Thylakoid
Membrane
Figure 6. Does PSII organization and mobility change in vivo? Schematic illustrating the dispersed and pseudocrystalline lattice arrangements of PSII 
(blue), which could potentially modulate an inverse relationship between local PSII concentration and mobility. Whether such a transition in PSII 
organization is physiologically relevant in vivo remains to be explored.
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sequential activities of superoxide dismutase and ascorbate per-
oxidase/catalase. The Mehler reaction can also generate a ΔpH 
across thylakoid membranes (Makino et al. 2002), although it is 
unclear how prevalent this reaction is in Chlamydomonas.
Pseudocyclic electron flow
Pseudocyclic electron flow (Fig. 7, #4) involves electron transfer to 
PSI-associated flavodiiron proteins (FLVs), likely through FDX 
(Sétif et al. 2020), which are thought to directly reduce O2 to H2O 
(Chaux et al. 2017). This reaction occurs in green algae, cyanobac-
teria, mosses, liverworts, and gymnosperms, but not in angio-
sperms, and is especially important immediately after dark 
acclimation when the Calvin-Benson cycle is not activated, and 
under fluctuating light conditions (Allahverdiyeva et al. 2013). 
A more detailed discussion of the flavodiiron proteins is given in 
the section below by Allahverdiyeva et al.
Plastoquinol terminal oxidoreductase
PTOX, associated with chlororespiration (Fig. 7, #5), uses electrons 
from the PQ pool to reduce O2 to H2O (Peltier et al. 2010). These 
electrons can come directly from PSII or from NADPH to the PQ 
pool via the NAD(P)H-PQ reductase; Type I NDH in plants and 
NDA2 in green algae (e.g. Chlamydomonas). Potential roles of 
PTOX include protection of cells during fluctuating light 
(Nawrocki et al. 2019) and PQ pool over-reduction, which can elicit 
PSII charge recombination and the generation of singlet O2. 
Chlamydomonas has two PTOX isoforms (PTOX1, PTOX2) 
(Houille-Vernes et al. 2011), with PTOX1 likely involved in regener-
ating oxidized PQ for phytoene desaturation (Foudree et al. 2012).
Chloroplast-to-mitochondria electron flow
The transfer of reducing equivalents between chloroplasts and 
mitochondria, here referred to as chloroplast-to-mitochondria 
electron flow (Fig. 7, #1a,1b), is also integral for controlling cellular 
energetics and redox conditions in the light, and for sustaining 
dark anabolic processes (Dang et al. 2014). Chloroplast–mitochon-
dria interactions were suggested in studies using inhibitors of mi-
tochondrial electron flow and mutants in chloroplast ATP 
synthase (Lemaire et al. 1988) and respiration (Cardol et al. 
2009). The integration between chloroplast and mitochondrial 
electron flow (He et al. 2023) helps coordinate the two pathways, 
balancing the production of reductant and ATP, regulating the 
metabolism of phosphate, nitrogen, and carbon compounds, 
and potentially managing accumulation of ROS to help sustain 
chloroplast electron transport and CO2 fixation. Major transloca-
tors for chloroplast-to-mitochondria electron flow have been pro-
posed to involve OMTs (malate/oxaloacetate, malate/aspartate) 
and TPTs. Plants and algae have multiple OMTs and TPTs, which 
are bidirectional transporters that function based on mass action 
and can move carbon and reductant out of chloroplasts (as these 
NDA2
FLVs
PTOXe−
Cyt
b6f
e−
Photons
H2O O2
O2
O2−
PSII 
(P680)
e−
PSI 
(P700)
NADP+
NAD(P)H
CBB cycle
+ NADPH
+ ATP
NAD(P)+
CO2
Triose-P
Triose-P
3-PGA
3-PGA
Pi +
+ Pi
TPT
Cytoplasm
Stroma
Thylakoid
membrane
5
1a
PQ
PC
FDX
ADP
+ PiATP
FNR
H2O2
H2O
O2
H2O
Malate
NADPH NADP+
OAA
OAA
Malate
OMT
OAA Malate
NADHNAD+
1b
PGR5
PGRL1
e−
e−
e−
2a 3
4
2b
Mitochondrion
H+
H+H+ H+
H+
Chloroplast
envelope
e− Photons
H+
?
Figure 7. Diagram of photosynthetic electron transport in Chlamydomonas, two major pathways for exporting fixed carbon and/or reductant from 
chloroplasts, and alternative pathways for electron flows (AEF). Linear photosynthetic electron transport involves excitation of reaction centers (PSI, 
PSII) and extraction of electrons from H2O by the PSII O2 evolving complex. Extracted electrons pass through PSII reaction centers, the plastoquinone 
(PQ) pool, Cytochrome b6f (Cyt), plastocyanin (PC) and to PSI where they are used to generate reduced ferredoxin (FDX) and NADPH; the ATP synthesized 
by the ATP synthase (fueled by proton gradient across thylakoid membranes) and the NADPH are used to drive CO2 fixation by the Calvin-Benson cycle. 
Major steps in these pathways are numbered in orange boxes. Fixed carbon and reductant are exported from chloroplasts by various shuttles; two 
major shuttles involve oxaloacetate/malate and triose-P/Pi exchange (OMT and TPT, respectively, 1a and 1b). Additionally, reducing electrons 
generated on the acceptor side of PSI or PSII can be routed through other AEF pathways (corresponding to 2 to 5) which include cyclic electron flow 
through both PGR5/PGRL1 (2a) and NDA2 (2b) pathways, the Mehler reaction (3) in which PSI-derived electrons are used to reduce O2 and the ROS 
generated can be converted to H2O through superoxide dismutase and catalase/ascorbate peroxidase, pseudocyclic electron flow (4) in which electrons 
can be used to reduce O2 to H2O through FLV flavodiiron proteins, and plastoquinol terminal oxidase (PTOX) catalyzed reduction of O2 (5) associated 
with the acceptor side of PSII. Major pathways associated with AEF are depicted with orange arrows while black arrows indicate linear electron flow.
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metabolites accumulate in the stroma), delivering them to other 
cellular compartments, including mitochondria. Reductant deliv-
ered by these transporters to mitochondria can serve as substrate 
for respiratory electron transport and reduction of O2 through cy-
tochrome oxidase or alternative oxidases (AOX), which are 
present in both plant and algal mitochondria. Inhibitor studies 
(myxothiazol, an inhibitor of the mitochondrial cytochrome bc1 
complex, and salicylhydroxamic acid, an inhibitor of the mito-
chondrial alternative oxidase) also suggest that the AOX pathway 
may generate some chemical bond energy/ATP that can support 
photosynthetic CO2 fixation through the formation of electro-
chemical potential (although AOX does not pump protons), but 
not as much as the cytochrome oxidase pathway (Peltier et al. 
2023). The AOX pathway is cyanide insensitive, enables continued 
operation of glycolysis and the TCA cycle under stress conditions 
when there is restricted electron flow through cytochrome oxi-
dase, balances C:N, ATP:ADP, NAD(P)H:ATP ratios, and limits 
hyper-reduction of the respiratory chain and accumulation of 
ROS (Rogov et al. 2014; Kaye et al. 2019).
While OMTs are thought to mainly shuttle reducing equiva-
lents between chloroplasts and other cellular compartments, 
TPTs may mainly export fixed carbon synthesized in chloroplasts 
(Huang et al. 2023). Chloroplast TPTs can transport triose-P [glyc-
eraldehyde 3-P (GAP), dihydroxyacetone-P (DHAP), and 3-phos-
phoglycerate (3-PGA)] in a counter exchange for cytosolic 
inorganic phosphate (Pi), which resupplies the chloroplast with 
Pi, allowing for continued photosynthetic electron flow and CO2 
fixation. TPTs are part of the pPT plastid translocator family 
that can serve as antiporters of phosphorylated C3, C5, or C6 com-
pounds with Pi (Flügge et al. 2003; Bockwoldt et al. 2019). In plants, 
trioses exported from chloroplasts are used to synthesize sucrose 
and other metabolites and fuel respiratory activity. Plants also 
harbor other pPT subfamilies, including those for glucose 6-P 
(GPTs) (Kammerer et al. 1998), xylulose-P (pentose-P) (Eicks et al. 
2002), and phosphoenolpyruvate (PEP) (PPTs) (Fischer et al. 
1997); PEP can be imported into C3 plant plastids and used as sub-
strate to fuel fatty acid biosynthesis or the shikimate pathway, 
and exported from C4 plant plastids (Streatfield et al. 1999; 
Prabhakar et al. 2010). In some plants, diminished TPT levels do 
not yield a strong phenotype because the loss of plastid TPT activ-
ity can be compensated for by accumulation of a transitory starch 
pool that can be rapidly degraded (Hausler et al. 1998, Walters 
et al. 2004) to products (e.g. hexose-P) exported from chloroplasts 
and used in other cellular compartments. Chlamydomonas TPT3 
was shown to likely be most active in routing triose-P out of chlor-
oplasts; tpt3 null mutants exhibited aberrant photosynthesis and 
highly reducing conditions in chloroplasts (Huang et al. 2023).
Contributions of AEF pathways
The relative importance of the different AEF pathways in generat-
ing the ATP needed to sustain CO2 fixation has been evaluated us-
ing electron transport inhibitors and mutants. Cyclic electron flow, 
pseudocyclic electron flow, and chloroplast-to-mitochondria elec-
tron flow activities were quantified in Chlamydomonas and shown 
to have compensatory activities, with each able to provide a large 
fraction of the energy needed to sustain photosynthesis (Peltier 
et al. 2023); the most energetically efficient pathway was 
chloroplast-to-mitochondria electron flow. However, using mu-
tants and pathway inhibitors to quantify individual pathway activ-
ities may not accurately reflect in vivo wild-type activities since a 
loss of any of these pathways in vivo may alter the contributions 
of the remaining pathways, and in some cases the inhibitors may 
not be completely penetrant.
Questions concerning integration of AEF activities
Photosynthetic electron flow and energy transfer are highly com-
plex processes (Fig. 7) and there is ongoing debate about the im-
portance of the different AEF pathways. Electron flow and 
energy transfer through these pathways largely occur through 
metabolic flux driven by mass action. This means that blocking 
one “road” can increase flux through another. Organisms can 
make adjustments by modifying gene expression or other regula-
tory phenomena. Some of the unknown questions include the 
following. 
1. Can we measure the precise contributions of each AEF path-
way in vivo without disrupting (mutants, inhibitors) the 
system?
2. What conditions govern the utilization of specific AEF activ-
ities and the proportion of each?
3. Which components of AEF complexes (e.g. cyclic electron 
flow) are involved in catalytic activity and regulation?
4. How do subunits of individual AEF complexes interact; what 
controls these interactions?
5. What components of AEF pathways are post-translationally 
modified; how do such modifications impact protein struc-
ture, interactions, and activities?
6. Why are there two different NDH pathways (Type I and II); do 
differences in these pathways reflect their ability to use dif-
ferent substrates?
7. What are the functions of the chloroplast OMTs and TPTs in 
driving chloroplast-to-mitochondria electron flow and what 
regulates their activities?
8. How are the activities of transporters of phosphorylated sug-
ars on the membranes of various cellular compartments inte-
grated with chloroplast sugar-P transport?
With an initial plunge into the waters of AEF and associated 
metabolite trafficking, we are just beginning to realize the depth 
of those waters.
Do flavodiiron proteins have multiple 
physiological roles in photosynthetic 
organisms?
(Written by Yagut Allahverdiyeva, Henna Mustila, 
and Lauri Nikkanen)
Flavodiiron proteins (FLVs) play a crucial role in some photosyn-
thetic organisms as an alternative electron transport pathway, al-
lowing growth in fluctuating light intensities by providing a valve 
for the safe dissipation of excess electrons in the photosynthetic 
electron transport chain. FLVs drive a four-electron reaction re-
ducing O2 to water downstream of PSI. This reaction is known as 
the Mehler-like reaction due to constituting a water–water cycle 
similarly to the Mehler reaction, or pseudocyclic electron trans-
port (Fig. 7 #4, and see above section by Grossman). FLVs are found 
in various photosynthetic organisms ranging from cyanobacteria 
to vascular plants, but are absent in red algae and angiosperms. 
Cyanobacterial Flv1 and Flv3 have structural and functional sim-
ilarities to FlvA and FlvB in algae and plants (Allahverdiyeva et al. 
2015a, 2015b, Ilík et al. 2017). The reason for the loss of FLVs from 
angiosperms during evolution remains elusive. However, it may 
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relate to increased efficiency of other electron sinks and protec-
tive mechanisms such nonphotochemical quenching, reactive 
oxygen species scavenging systems, and cyclic electron transport, 
which would have made retention of energy-wasting FLVs redun-
dant or even disadvantageous (for discussion on this, see e.g. 
Alboresi et al. 2019b).
In the reference cyanobacterium Synechocystis (Synechocystis sp. 
PCC6803), the Flv1/Flv3 hetero-oligomer catalyzes strong but tran-
sient O2 photoreduction in the first minute of illumination or dur-
ing increased light intensity under ambient CO2 (Santana- 
Sánchez et al. 2019), providing essential electron sink capacity dur-
ing the initial phase of photosynthetic induction before full activa-
tion of the Calvin–Benson cycle (Fig. 8A). Accordingly, flv mutants 
(deficient in Flv1 or Flv3 or homologs FlvA and FlvB) demonstrate 
strong growth retardation under fluctuating light, where low back-
ground light is repeatedly interrupted by high-light pulses, causing 
strong acceptor-side limitation of PSI (Allahverdiyeva et al. 2013; 
Gerotto et al. 2016; Chaux et al. 2017; Jokel et al. 2018). 
Synechocystis Flv mutants show a less pronounced phenotype 
under milder fluctuating light conditions (higher background light) 
at pH 8.2, and no growth phenotype was observed at pH 7.5 (Mustila 
et al. 2016), suggesting an external pH-dependent effect on FLVs 
and growth that remains poorly understood.
In addition to Flv1 and Flv3, some β-cyanobacteria possess Flv2 
and Flv4 proteins (Helman et al. 2003). These proteins also partic-
ipate in Mehler-like reactions downstream of PSI, but with differ-
ent kinetics, catalyzing O2 photoreduction that persists beyond 
the initial phases of photosynthetic induction (Santana-Sánchez 
et al. 2019; Fig. 8A). Unlike Flv1 and Flv3, Flv2 and Flv4 are not cru-
cial during sudden increases in light intensity. In Synechocystis, 
Flv2 and Flv4 show strong downregulation under low light, high 
CO2 and high pH conditions (Eisenhut et al. 2012; Santana- 
Sánchez et al. 2019). Deletion of Flv2 and Flv4 alters the phenotype 
under high light, and overexpression of flv4-2 operon provides re-
sistance to high light (Bersanini et al. 2014). Notably, the presence 
of these proteins within an operon alongside Sll0218 suggests that 
some phenotypic changes could be attributed to the role of this 
small protein in PSII assembly and stabilization (Bersanini et al. 
2017).
Heterocyst-forming N2-fixing cyanobacteria, such as Anabaena 
(Anabaena sp. PCC 7120), possess six FLVs (Fig. 8C). Flv1A and 
Flv3A are specific to vegetative cells and are crucial for survival 
under FL conditions. Interestingly, deletion of Flv3A affects N2 fix-
ation and H2 metabolism in heterocysts, specialized N2 fixing 
cells, via downregulation of uptake hydrogenase, enabling H2 pro-
duction even under oxic conditions (Santana-Sánchez et al. 2023). 
Figure 8. Function and regulation of FLVs in Synechocystis and Anabaena. A) In Synechocystis, Flv1/Flv3 and Flv2/Flv4 hetero-oligomers function in 
light-driven O2 photoreduction and their expression level is dependent on carbon availability. In addition, Flv3/Flv3 homo-oligomer catalyzes a distinct, 
unspecified reaction than the O2 photoreduction. B) The reversible association of different Flv1/3 and Flv2/4 hetero-oligomers with the thylakoid 
membrane of Synechocystis is controlled by the extent of the trans-thylakoid proton gradient. The + symbols refer to positively charged protons. See 
text for details. C) In filamentous Anabaena, Flv1A/Flv3A function in vegetative cells and Flv3B/Flv3B in heterocysts in O2 photoreduction, the latter 
configuration maintaining microoxic conditions to protect the N2-fixing enzyme nitrogenase in the light. The roles of Flv2/Flv4 and Flv1B/Flv1B in 
Anabaena are yet to be elucidated.
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Investigations into the roles of Flv2 and Flv4 in Anabaena are 
yet to be conducted, although their expression pattern under 
different conditions suggests a function similar to that in 
Synechocystis. Flv1B and Flv3B are specific to heterocysts 
(Ermakova et al. 2013, 2014). Unlike other cyanobacterial FLVs, 
Flv3B can function as a homo-oligomer in the Mehler-like reac-
tion, maintaining microoxic conditions under light. Importantly, 
while Flv1B is not involved in the Mehler-like reaction, its deletion 
strongly impacts N2 fixation, indicating a role in N2 metabolism 
(Ermakova et al. 2014).
What are the functional oligomeric states of FLVs?
All FLVs contain an N-terminal metallo-β-lactamase-like domain 
harboring an O2 and/or NO reducing nonheme diiron center and a 
C-terminal flavodoxin-like (FMN) domain (Vicente et al. 2008). 
Oxygenic photosynthetic organisms contain an additional 
NAD(P)H:flavin reductase-like domain fused in the C-terminus. 
In vitro studies and modeling suggested that FLVs function in a 
head-to-tail homo-oligomeric configuration (Vicente et al. 2008) 
enabling inter-subunit electron transfer, between Fe–Fe of one 
monomer and FMN of the other. In Synechocystis, an in vivo 
hetero-oligomeric configuration (Flv1/3 and Flv2/4) is important 
for O2 photoreduction: Synechocystis strains lacking Flv1 but 
overexpressing Flv3 (thus Flv3/3 configuration) or lacking Flv3 
but overexpressing Flv1 (Flv1/1) did not demonstrate O2 photore-
duction (Mustila et al. 2016). However, having overexpressed 
homo-oligomers of Flv3 or Flv1 partly rescued the phenotype 
under FL, suggesting that homo-oligomers can function in un-
known metabolic pathways and play a role in photoprotection 
that is independent of O2 photoreduction. Moreover, in 
Anabaena, the deletion of Flv1A did not result in total impairment 
of the Mehler-like reaction in air-level CO2 conditions where Flv2 
and Flv4 are expressed (Santana-Sánchez et al. 2023). This sug-
gests that Flv3A can catalyze O2 photoreduction in an Flv2/ 
4-dependent manner either as a homo-oligomer interacting with 
Flv2/4 or by forming oligomers with Flv2 or Flv4.
Can FLVs in photosynthetic organisms catalyze 
NO reduction?
FLVs can reduce NO into N2O in anaerobic bacteria (Vicente et al. 
2008) and in Chlamydomonas reinhardtii (Burlacot et al. 2020). We 
cannot exclude a similar function in cyanobacteria, although 
the efficiency of this reaction as a sink is disputable. However, it 
may affect cell metabolism via signaling, and could be aligned 
with alternative electron donors. Recent data show that 
Synechocytsis Fed1, the most abundant of 11 ferredoxins in this 
cyanobacterium, can interact with and act as a donor to the 
Mehler-like reaction catalyzed by Flv1/3 and Flv2/4 (Nikkanen 
et al. 2020, 2023; Sétif et al. 2020). However, in vitro studies dem-
onstrated that NAD(P)H is a donor for recombinant Flv1, Flv3, or 
Flv4 homo-oligomers (Vicente et al. 2008; Shimakawa et al. 2015; 
Brown et al. 2019), albeit with relatively low electron transfer 
rates. Interestingly, a protein–protein interaction was detected be-
tween Flv3 (but not Flv1) and the large isoform of FNR (FNRL) 
(Nikkanen et al. 2023). Flv3/FNRL interactions could occur to facil-
itate electron transfer from NADPH to Flv3 homo-oligomers, func-
tioning in a pathway to reduce an unknown acceptor, such as NO. 
Flv1-independent activity of Flv3 is also supported by 6-fold high-
er protein amount of Flv3 in comparison to Flv1 (Jackson et al. 
2023). However, the physiological relevance of these interactions 
and Flv3-homo-oligomer-dependent electron transfer pathways 
remain unresolved.
How is FLV activity regulated?
The Mehler-like reaction displays distinct kinetics where Flv1/ 
3-dependent O2 photoreduction is transiently induced and then 
inactivated during the first minute of illumination. This strongly 
suggests that FLV activity is subject to post-translational regula-
tion (Alboresi et al. 2019a). Recently, we proposed a model for 
pH-dependent regulation of FLV hetero-oligomer activity 
(Nikkanen et al. 2023), where alkalization of the cytosol during 
photosynthetic induction results in Flv1/3 hetero-oligomers being 
repelled from the thylakoid membrane due to their surface charge 
becoming more negative and their activity being thus decreased 
(Fig. 8B). Flv2/4 remains associated with the thylakoids longer 
after photosynthetic induction (i.e. in a more alkaline cytosol) 
due to large positively charged patches in Flv4. While this model 
explains many features of the observed kinetics of FLV activity, 
FLVs are likely subject to additional layers of post-translational 
regulation. The fact that we see a transient reduction of ferredox-
in at all during dark-to-light transitions suggests that Flv1/3 
hetero-oligomers are not yet fully primed in darkness, despite cy-
tosolic pH being neutral and FLV hetero-oligomers being associ-
ated with the thylakoids. Therefore, it is likely that an additional 
unknown signal is required for the full light-dependent activation 
of Flv1/3 hetero-oligomers. Indeed, FLVs have been suggested to 
be redox-regulated via conserved cysteine residues, some of 
which in Flv1 and Flv3 are light-dependently reduced (Guo et al. 
2014). It remains to be elucidated if thiol modulation or other post- 
translational modifications, such as phosphorylation, play a role 
in regulating FLV activity.
An additional question concerns how FLV activity is regulated 
in low versus high CO2 conditions. In Synechocystis, in high CO2, 
although Flv1 and Flv3 are downregulated, and Flv2 and Flv4 are 
not expressed (Santana-Sánchez et al. 2019), FLV-dependent O2 
photoreduction activity is sustained at a higher level than in low 
CO2. This suggests a regulatory mechanism that connects FLV ac-
tivity to carbon metabolism and possibly to carbon concentrating 
mechanisms.
How is photosynthesis regulated during 
light fluctuations?
(Written by Adrien Burlacot)
Photochemical reactions happen extremely fast (on the picosec-
ond timescale), but the redox reactions involved in the transport 
of electrons are orders of magnitude slower, and the metabolic re-
actions of carbon metabolism are even slower. This timescale dis-
crepancy makes meeting metabolic energy requirements with 
light-energy-driven PS activity challenging to attain when light in-
tensity is variable. In extreme cases of light fluctuation, photosyn-
thetic electron generation can overflow electron transport or 
metabolic capacity, leading to PS degradation (Sejima et al. 
2014) and potentially cell death. In many natural contexts, light 
rapidly fluctuates between a sub-saturating (low light, LL) to an 
over-saturating level (high light, HL) (Fig. 9A), and cells must 
cope with either excessive or insufficient amounts of energy and 
quickly switch between those states as light changes. The electron 
transport chain is a central hub of regulation in this context. 
Despite decades of investigations, how photosynthesis is dynam-
ically controlled to provide chemical energy without becoming 
harmful during light fluctuations remains a thriving area of 
research (Morales and Kaiser 2020). This question has recently 
regained traction with groups showing that tuning the dynamics 
of light energy management can improve the productivity of 
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photosynthetic organisms (Kromdijk et al. 2016; De Souza et al. 
2022; Perin et al. 2023). Previous sections provided an overview 
of alternative electron flow pathways (Grossman) and the role 
of the pathway mediated by FLVs in some organisms 
(Allahverdiyeva et al.). Here, I discuss questions related to how or-
ganisms regulate the dynamics of which pathways and mecha-
nisms are employed under fluctuating light conditions in 
different environments.
Many dynamic regulatory mechanisms of photosynthetic elec-
tron generation and management are triggered by low lumenal 
pH, which modulates electron transport at the level of the cyto-
chrome b6f (Malone et al. 2021) and activates nonphotochemical 
quenching (NPQ) of excess absorbed light energy. Since the dis-
covery of the role of thylakoid protein phosphorylation in 
response to 5 min LL/1 min HL treatment (Fig. 9B) (Tikkanen 
et al. 2010), similar artificial light patterns have been extensively 
used to identify proteins involved in the adjustment of the ener-
getic status of the electron transport chain when light fluctuates. 
In model species of angiosperms, mosses, microalgae, and cyano-
bacteria (Fig. 9B) those include: (i) proton sensing proteins in-
volved in NPQ which limit photochemistry and protect PS during 
HL (Cantrell and Peers 2017; Roach 2020; Steen et al. 2022; Niu 
et al. 2023); (ii) ion exchange across the thylakoid membrane 
mediated by VCCN1 and KEA3, which balances the proton motive 
force and controls NPQ and electron flow during the few minutes 
following a light change (Armbruster et al. 2016; Herdean et al. 
2016), (iii) the plastid terminal oxidase (PTOX), which contributes 
to the redox poise of the thylakoid during the LL/dark phase of 
O2
CO2
O2
Redox
shuttling
K
E
A
3
V
C
C
N
1
N
P
Q
LH
C
II
PSII
STN7, 
STN8, 
STT7
A C
June 09 2021June 10 2021
June 16 2021 June 11 2021
0
500
1,000
1,500
2,000
2,500
Li
gh
t
in
te
ns
ity
(µ
m
ol
ph
ot
on
m
−2
s−
1 )
Li
gh
t
in
te
ns
ity
(µ
m
ol
ph
ot
on
m
−2
s−
1 )
0
0 6 12 18 24
500
1,000
1,500
2,000
2,500
Time (h)
0 6 12 18 24
Time (h)
B
Functional domain of
importance
(molecular mechanisms)
Prediction of importance
under complex light patterns
Average
Maximum
Minimum
Periodicity
In
te
ns
ity
...Others
Time
Cytosol
Chloroplast 
stroma
Lumen
LH
C
II
P
K+
H+
Cl−
PSI
H+
H+H+
H+
H+
e− e−
e−
e−
PTOX
FLVsPGR5,
PGRL1
Cyt
b6f
Figure 9. Acclimation mechanisms to light fluctuations. A) Light fluctuations generated by diurnal cycle and cloud coverage on top of a field canopy in 
June 2021. Data shown are light intensity of the photosynthetically active radiations throughout the time of the day from the Surface radiation Budget 
Network (SurfRAD, NOAA) at the Bondville Station, Illinois (40°N latitude; 88°W longitude). B) Main molecular mechanisms involved in tuning 
photosynthetic electron transport during artificial light fluctuations. The lumenal pH controlled by ion transport (mediated by KEA3, VCCN1, and the 
ATPase) and alternative electron transport (mediated by PGR5/L1 and FLVs), activates NPQ mechanisms and controls the photosynthetic electron flow 
at the cytochrome b6f (Cyt b6f ) level. Phosphorylation of LHCII, which dynamically relocates LHCII to PSI, and redox exchange between the chloroplast 
and the rest of the cell are also involved in the dynamic regulation of photosynthetic electron transport. Shown in blue arrows are photosynthetic 
electron transport pathways. C) Example of characteristics of light fluctuations that can be explored to define the domain of importance of molecular 
mechanisms involved. The definition of such domains might help in predicting the set of mechanisms relevant to survive in a more complex light 
pattern.
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fluctuation (Nawrocki et al. 2019), (iv) cyclic electron flow con-
trolled by PGR5 and PGRL1 proteins which tunes lumenal pH 
and electron flow during the first minutes of HL (Yamamoto 
et al. 2016; Yamamoto and Shikanai 2018; Storti et al. 2019), (v) 
FLVs, which dissipate the overflow of electrons and contribute 
to the low lumenal pH upon HL exposure within a few seconds 
by reducing O2 to water (Allahverdiyeva et al. 2013; Gerotto et al. 
2016; Chaux et al. 2017; Jokel et al. 2018), and (vi) redox shuttles 
which rebalance the redox poise between the organelles 
(Yokochi et al. 2021).
Despite this extensive amount of knowledge, the complete set 
of proteins involved during light fluctuations is likely just begin-
ning to be revealed. Indeed, acclimation mechanisms to HL or 
LL involve (i) posttranslational regulation of protein activity 
(Allorent et al. 2013) (on a timescale of milliseconds to tens of 
minutes), (ii) translation of mRNA (Durnford et al. 2003) (on a 
timescale of minutes), (iii) changes of mRNA levels (Mettler et al. 
2014), (iv) structural changes of organellar membranes 
(Kirchhoff 2014b) or (v) leaf orientation or morphology (Feng 
et al. 2019) (on a timescale of minutes to hours/days). Hence, 
the preceding time in LL or in HL, the intensities of LL and HL, 
the shape of the light transient as well as how frequent light is 
shifting likely have a major impact on the identity and role of 
mechanisms involved in acclimation. The use of just one light 
fluctuation regime becomes thus limited both for discovering 
new proteins involved and for identifying domains where known 
proteins are important.
How do different light fluctuation characteristics change 
which mechanisms are involved in photosynthesis regulation? 
Reductionist approaches hold great promises in that regard. One 
of them, introduced by Nedbal and Březina (2002) and recently 
used on mutated strains by Shimakawa and Miyake (2018); 
Steen et al. (2022), and Niu et al. (2023) involves assessing the re-
sponse of mutants impaired in photosynthetic regulatory proc-
esses to artificial periodic illumination (square-like or sine-like). 
Varying the periodicities used over short-term episodes of light 
fluctuations allows us to define the periodicity domain of photo-
synthetic regulatory processes, which are periodicity limits below 
or above which a regulatory mechanism cannot dynamically re-
spond to a periodic fluctuation. Defining the periodicity domain 
of an acclimatory mechanism might allow predicting its impor-
tance during a more complex light fluctuation harboring a wide 
set of fluctuation periodicities. Note here that while this method 
is common in engineering sciences and relates to frequency- 
domain analysis, I refer here to the periodicity domain for nota-
tion simplicity. This method has allowed the unraveling of perio-
dicity domains of proteins involved in NPQ: (i) in the green algae 
Chlamydomonas, NPQ magnitude and dynamics depend more 
on the light-harvesting complex stress-related 3 protein 
(LHCSR3) for a 1 min periodicity than for a 10 min periodicity 
(Steen et al. 2022), and (ii) in the angiosperm Arabidopsis, NPQ 
does not respond to dynamic light changes below a periodicity 
of 10 s while for periodicities of 10 to 60 s, the protein PsbS is crit-
ical for NPQ response; for longer periodicities, both PsbS and 
changes in the xanthophyll cycle contribute to the NPQ response 
(Niu et al. 2023). However, while defining periodicity limits of ac-
climatory processes can prove important, it remains limited to 
short-term episodes of light fluctuation and misses long-term ac-
climatory mechanisms. Another approach consists of growing/ 
acclimating the photosynthetic organism under different patterns 
of light fluctuation and measuring how photosynthetic parame-
ters are affected by the growth/acclimation conditions. Such an 
approach can reveal emergent phenomena in new mutants 
(Peng et al. 2020) and has recently revealed that (i) the average 
growth light intensity affects the quantum efficiency of PSII and 
the dynamics of ion transport across the thylakoids upon HL 
while, (ii) the variability of growth light changes the levels and 
de-epoxidation state of the xanthophyll-cycle pigments violaxan-
thin, antheraxanthin, and zeaxanthin (von Bismarck et al. 2023). 
Future work combining those two approaches with other key 
characteristics of light fluctuations will be important to under-
stand how the variability of light influences the mechanisms in-
volved in acclimation to light.
What happens in a natural light environment? While fluctua-
tions of light intensity in nature can be complex (Fig. 9A), reduc-
tionist methods like the ones discussed above are powerful and 
should allow, when used systematically, the prediction of the 
identity and activity of acclimatory processes involved depending 
on key characteristics of light fluctuations (Fig. 9C). The perio-
dicity of light fluctuation is a clear characteristic of natural rele-
vance, as under a forest canopy, the movement of leaves forms 
gaps that let high-intensity sunlight pass typically during less 
than 1.6s (Pearcy 1990) while, at the top of the canopy, HL expo-
sure typically lasts for a few minutes (Smith and Berry 2013). 
Such periodicity variability might even be larger in aquatic envi-
ronments where rippling caustic effects of the water surface fluc-
tuate light with periodicities of less than 100 ms (Schubert et al. 
2001). While other light fluctuation characteristics influencing 
light acclimation mechanisms like average, maximal, and mini-
mal light intensity have been explored to some extent 
(Allahverdiyeva et al. 2013; Jokel et al. 2018; von Bismarck et al. 
2023); identifying a subset of key light characteristics that de-
scribes natural light fluctuations in different environment and 
quantifying their range remains to be done. An interesting possi-
bility here is to apply signal decomposition methods currently 
used to study the output of photovoltaic panels in electrical engi-
neering (Meyers and Boyd 2023) to define what are the most rele-
vant light intensity characteristics present in different natural 
light environments.
Importantly, the presence or the absence of proteins regulating 
photosynthesis in different species can vary widely and this diver-
sity and its implication are critical to study. For example, while 
FLVs are required for acclimation to 1min HL/5min LL fluctuations 
in many phototrophs, they are absent in angiosperms (Yamamoto 
et al. 2016), the latter most likely relying on other mechanisms to 
cope with sudden HL transients. The development of knowledge 
on the diversity of mechanisms across species (Sello et al. 2019), 
together with modeling the potential interaction of different 
mechanisms (Li et al. 2021a, 2021b), might allow molecular breed-
ing of plants for adaptation to their local light environment in con-
ditions where acclimation to light limits growth (Perin et al. 2019).
How is photosynthesis regulated during light fluctuations? The 
development of new tools (Cruz et al. 2016) and methods (Nedbal 
and Lazár 2021) to systematically look at different aspects of this 
question will bring new perspectives to the complex regulations of 
photosynthesis under light fluctuations. And the answers will 
most likely depend on which characteristics of light fluctuation 
are investigated.
How can photorespiration impact stomatal 
opening?
(Written by Michael Hodges)
Photosynthetic CO2 assimilation in embryophytes requires the 
uptake of atmospheric CO2 into the leaf. This is achieved by sto-
mata, specialized epidermal structures composed of two guard 
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cells. They possess partially overlapping signal transduction net-
works controlling stomatal pore size by modulating guard cell tur-
gor pressure in response to abiotic and biotic cues including light, 
CO2, water potential, temperature, ozone, and pathogens. In this 
way, plants balance CO2 uptake and water loss with respect to 
fluctuating environmental conditions. Stomata are a target to be 
considered when designing future-proofed crops (Nguyen et al. 
2023), therefore a complete understanding of how stomatal open-
ing and closure are regulated is important. Although there is 
abundant knowledge concerning guard cell signaling networks 
and the H + -pumps, H + -coupled ion transporters, and ion chan-
nels involved in controlling stomatal movements (Saito and 
Uozumi 2019), how plant metabolism plays a role is less clear 
(Lemonnier and Lawson 2023). Mitochondrial activity (Nunes- 
Nesi et al. 2007) and cytosolic ATP (Lim et al. 2022) appear to be 
important for stomatal opening. Low blue light perceived by 
PHOT1/2 photoreceptors is involved in early morning guard cell 
starch degradation required for stomatal opening (Horrer et al. 
2016). Photosynthesis has been shown to promote stomatal open-
ing but its relative importance and cellular location (mesophyll 
cells versus guard cells) are still open to debate (Lemonnier and 
Lawson 2023). Recently, a role for photorespiration in stomatal 
functioning was suggested from experiments using photorespira-
tory mutants (Eisenhut et al. 2017; Flügel et al. 2017; Duminil 
2019). If this is so, then how do photorespiratory cycle activity 
and/or photorespiratory-associated metabolites interfere with 
processes and signaling pathways that control stomatal open-
ing/closure?
So what is photorespiration and which photorespiratory me-
tabolites have the potential to influence stomatal opening/clo-
sure? Photorespiration is an essential metabolic process taking 
place in the light. It begins in chloroplasts with the oxygenase ac-
tivity of Rubisco producing 2-phosphoglycolate (2-PG), which can 
inhibit the Calvin–Benson–Bassham cycle enzymes triose- 
phosphate isomerase and sedoheptulose 1,7-bisphosphate 
phosphatase as well as glycolytic phosphofructokinase (Flügel 
et al. 2017 and references therein). The photorespiratory cycle 
spans four cell compartments (chloroplasts, peroxisomes, mito-
chondria, cytosol) and requires eight enzymes to transform toxic 
2-PG into useful 3-phosphoglycerate. To accomplish this, photo-
respiration leads to the production of H2O2 and NADH while liber-
ating CO2 and ammonia during serine formation from glycine in 
mitochondria (reviewed in Eisenhut et al. 2019).
As early as the 1960s, short-term pharmacological studies us-
ing photorespiratory enzyme inhibitors showed a correlation be-
tween photorespiration and stomatal opening in the light. The 
addition of hydroxysulfonate (an inhibitor of glycolate oxidase, a 
peroxisomal enzyme producing H2O2 and glyoxylate from glyco-
late) to tobacco leaf disks (Zelitch and Walker 1964) and aminoa-
cetonitrile (an inhibitor of glycine decarboxylase, a mitochondrial 
complex producing CO2) to rice leaves (Cho et al. 1987) provoked 
stomatal closure in the light. An association between photorespi-
ration and stomatal opening/closure was further suggested be-
cause the inhibitory effect of aminoacetonitrile did not occur in 
detached rice leaves under low photorespiratory O2 air conditions 
and aminoacetonitrile application to maize (Zea mays) leaves (a C4 
plant with low photorespiration) did not alter transpiration (Cho 
et al. 1987).
A renewed interest in photorespiration and stomata opening/ 
closure has come from recent observations made using 
Arabidopsis photorespiration mutants (Fig. 10). Compared to wild- 
type (WT) plants, mutants lacking either phosphoglycolate phos-
phatase (pglp1), or serine hydroxymethyltransferase (shm1), or 
glycerate kinase (glyk1) showed higher and lower transpiration 
rates at high CO2 (10,000 ppm) and in ambient air, respectively 
(Eisenhut et al. 2017). This was later partially confirmed by direct 
stomatal aperture measurements using epidermal peels of pglp1 
leaves and light microscopy (Duminil 2019). Arabidopsis PGLP1 
antisense repression also led to lower stomatal conductance (gs) 
and transpiration rates when plants were transferred from high 
Figure 10. Potential connections between photorespiration and metabolic processes relevant to stomatal opening/closure. In the light, Rubisco is 
involved in both photosynthetic CO2 and photorespiratory O2 assimilation. Photorespiration produces a number of metabolites that have the potential 
to impact stomatal opening and closure. 2-PG can reduce starch biosynthesis and therefore blue light/PHOT-induced malate from starch degradation 
and thus limit early morning stomatal opening. 2-PG might also inhibit glycolysis and malate biosynthesis required for stomatal opening while 
photorespiratory CO2 and 3-phosphoglycerate (3-PGA) could be used to produce malate via PEP carboxylase (PEPC) activity for stomatal opening. 
Photorespiration also produces H2O2 and serine, and both could be linked to stomatal closure either as a signaling molecule or via the production of 
O-acetylserine required for sulfate-induced ABA biosynthesis, respectively. Photorespiration also negatively impacts photosynthetic activity thus 
reducing the capacity to produce starch, sugars, and ATP that are important for stomatal opening.
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to ambient CO2 (Flügel et al. 2017). On the other hand, PGLP1 over-
expression gave higher gs values and transpiration rates in 40% O2 
air; a high-photorespiration condition that increased leaf 2-PG 
amounts (Flügel et al. 2017). Lines overexpressing PGLP1 also 
maintained high gs levels at an elevated temperature and after 
drought stress, conditions associated with increased photorespi-
ration (Timm et al. 2019). Greenhouse-grown tobacco overex-
pressing glycine decarboxylase (GDC)-H subunit also exhibited 
higher gs although this was not observed in the field 
(López-Calcagno et al. 2019). By analyzing epidermal peels of com-
plemented Arabidopsis shm1 lines, SHM activity was found to im-
pact salt- and ABA-induced stomatal closure (Liu et al. 2019).
So how can photorespiration impact stomatal opening/closing? 
There are a number of possible mechanisms that require investi-
gation (Fig. 10). 
• Transcriptional regulation could be involved since several 
photorespiratory mutants have deregulated genes involved 
in stomatal closure including ABSCISIC ACID INSENSITIVE 
1/2 (ABI1/2) (ABA biosynthesis), and RESPIRATORY BURST 
OXIDASE HOMOLOGS D/F (RBOHD/F) (reactive-oxygen produc-
tion) and blue-light-induced stomatal opening (PHOT1/2) 
(Eisenhut et al. 2017). The signals and mechanisms involved 
are not yet known.
• Photorespiration could act by indirectly or directly impacting 
photosynthetic electron transfer and as a consequence ATP 
biosynthesis, which is required for stomatal opening. 
Photosynthetic processes have been shown to be affected by 
reduced photorespiratory cycle activity in mutants, including 
an attenuation of photosynthetic CO2 assimilation, modifica-
tion of PSII redox potential, and an increase in energy dissipa-
tion mechanisms (Timm et al. 2012; Dellero et al. 2015, 2016; 
Messant et al. 2018). Indeed, a correlation between gs and pho-
tosynthesis exists (Nunes-Nesi et al. 2007; Lu et al. 2014).
• Changes in photorespiratory activity could also provoke met-
abolic regulation by interacting with and perhaps modulating 
metabolic pathways relevant to stomatal movements includ-
ing the biosynthesis of starch, malate, and ABA.
• Photorespiration could impact blue-light-induced stomatal 
opening at the beginning of the day. It has been proposed 
that enzyme inhibition by 2-PG is a control loop adjusting sto-
matal conductance to photosynthesis-related C-fluxes by al-
tering C-allocation between RuBP regeneration and starch 
synthesis (Flügel et al. 2017). A correlation between PGLP ac-
tivities, 2-PG levels, gs, and transpiration rates in PGLP trans-
genic lines (Eisenhut et al. 2017; Flügel et al. 2017; Timm et al. 
2019) and reduced end-of-day leaf starch levels in photores-
piratory mutants (Timm et al. 2012) support this hypothesis.
• Guard cell osmoregulation requires starch breakdown, triose- 
phosphate export from chloroplasts, and cytosolic PEP carbox-
ylase (PEPC) activity for glycolytic malate production (Gehlen 
et al. 1996). Photorespiration produces 3-PGA and liberates 
CO2 and therefore this could fuel glycolysis and PEPC activity 
to promote guard cell malate biosynthesis and stomatal open-
ing. On the other hand, a build-up of photorespiratory 2-PG 
might inhibit glycolysis and favor stomatal closure.
• Sulfate-induced stomatal closure upon drought is associated 
with guard cell-specific ABA biosynthesis (Batool et al. 2018). 
In photosynthetic tissues, photorespiration is the predomi-
nant serine biosynthesis route with 23% to 41% of photores-
piratory serine leaving the cycle (Fu et al. 2023) to serve 
metabolic processes including sulfur-assimilation and 
cysteine biosynthesis (Abadie and Tcherkez 2019). In this con-
text, photorespiratory serine could produce o-acetyl serine re-
quired for sulfate-associated cysteine biosynthesis and 
subsequent ABA production.
So is the photorespiratory cycle a good target to improve sto-
matal opening/closure traits in the field? Chloroplast photorespir-
atory by-pass plants with improved photosynthesis might be 
expected to impact stomatal opening. That said, no significant 
changes were found in Arabidopsis (Maier et al. 2012) or tobacco 
(South et al. 2019), and only rice showed a higher gs (Nayak 
et al. 2022). Not all photorespiratory mutants exhibit altered gs 
or transpiration when transferred from high CO2 to air (Dellero 
et al. 2015, 2016; Eisenhut et al. 2017) although antisense rice 
GOX showed a correlation between GOX activity and gs (Lu et al. 
2014). Finally, reduction of end-of-day starch in hpr1 leaves 
(Timm et al. 2012) was not associated with altered transpiration 
in the light (Eisenhut et al. 2017).
Many questions still remain unanswered such as how, where, 
and when does photorespiration impact stomatal opening/clos-
ing? It will be challenging but essential to answer these questions 
in the context of manipulating photorespiration to improve pho-
tosynthesis and plant yield since unwanted stomatal phenotypes 
could negatively impact water-use efficiency. Future directions to 
answer these questions should include determining stomatal ki-
netic responses to environmental cues in controlled atmospheric 
conditions differing in CO2/O2 levels to modulate photorespiration 
in attached leaves. Other important avenues of investigation in-
clude determining whether photorespiration in guard cells, mes-
ophyll cells, or both cell types is important using cell-specific 
expression lines, and identifying how photorespiration impacts 
metabolites involved in stomatal movements using single-cell 
metabolomics associated with stable-isotope labeling.
What adaptive changes make an enzyme 
suitable for the C4 carbon concentrating 
pathway, and how do critical and 
lineage-specific amino acid substitutions 
affect plant physiology?
(Written by Clarisa E. Alvarez, Veronica 
G. Maurino, and Marcos A. Tronconi)
C4 proteins have accumulated different types of 
adaptive changes
Photorespiration evolved as a selective response to Rubisco 
promiscuity, as a high-energy pathway to remove 2-phosphogly-
colate, an unwanted reaction product. In C3 plants, photorespira-
tion negatively affects fitness under conditions that reduce the 
relative concentration of CO2, such as high temperature, drought, 
and salinity. Some land plants have evolved a biochemical CO2 
pump known as the C4 carbon concentrating pathway or C4 path-
way. In the C4 pathway, CO2 is prefixed to form a C4 acid, which is 
then decarboxylated in the vicinity of Rubisco. These reactions are 
usually spatially separated between two cell types (Fig. 11A) 
(Maier et al. 2011; Drincovich et al. 2016). By increasing the CO2 
concentration around Rubisco, C4 plants gain a beneficial reduc-
tion in photorespiratory flux. The evolution of the C4 pathway re-
sulted in the coordinated acquisition of new anatomical and 
biochemical features (Sage and Zhu 2011; Alvarez and Maurino 
2023). It is widely accepted that anatomical C4 traits were ac-
quired prior to C4 biochemistry (Christin and Osborne 2014). On 
this basis, C4 pathway genes were formed by co-opting copies of 
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duplicated genes that already existed in C3 species (Christin et al. 
2013). In most cases, the same gene lineage has been selected re-
peatedly by independent C4 origins. The bias towards co-opted 
genes may be related to fewer key genetic changes in their regula-
tory and coding regions, which need to accumulate rapidly to ac-
quire the C4 function (Copley 2020).
Some of the adaptive amino acid changes are critical for C4 
function: if one of these is mutated back to the amino acid present 
in the C3 version of the enzyme, there is a loss of crucial C4 bio-
chemical properties (kinetic, regulatory, and/or structural). In ad-
dition, other adaptive changes have occurred to optimize the C4 
function, either (i) by counteracting perturbations in protein prop-
erties caused by previous critical C4 changes, such as a reduction 
in stability of the protein structure, or (ii) by adapting the enzyme 
activity to the new physiological environment that accompanies 
C4 activities. Candidates for critical adaptive amino acid changes 
can be identified through an analysis of protein sequences from 
different C4 lineages in the form of strictly differentially substi-
tuted positions (Alvarez et al. 2019; Hüdig et al. 2022). A strictly dif-
ferentially substituted position is one where the protein 
sequences in the C4 species contain an identical amino acid, while 
a different amino acid is shared by all C3 sequences. The data ob-
tained so far indicate that critical adaptive changes for C4 innova-
tions have evolved with a high degree of convergence between 
distant C4 lineages, as in the case of PEPC (Christin et al. 2007; 
Rosnow et al. 2014; Lyu et al. 2021), or restricted to genetically 
close C4 lineages, as in the case of NADP-ME (Alvarez et al. 2019; 
Alvarez and Maurino 2023) and NAD-ME (Tronconi et al. 2020; 
Hüdig et al. 2022), demonstrating that alternative solutions to crit-
ical C4 requirements exist. Such sequence analyses also reveal a 
large number of molecular adaptations that are lineage-specific 
(Fig. 11, B and C).
Innovation is easy, optimization is complicated: 
how can we analyze the impact of adaptive 
changes of C4 proteins?
C4 innovations can be achieved by a rather small number of crit-
ical changes, some of which evolved divergently while others 
appear to have occurred convergently across C4 lineages. In this 
regard, when comparing the differentially substituted residues 
between the C4-PEPC from Poaceae, Chenopodiaceae, and 
Asteraceae, which are genetically very distant, only one residue 
underwent an identical substitution in all three lineages. This 
substitution causes the C4-PEPC to acquire affinity values for sub-
strates similar to those of non-C4 isoforms. This suggests that only 
minimal changes to the C3 ortholog may be required to perform 
the specialized C4 function. However, the presence of a large num-
ber of lineage-specific amino acid substitutions in C4 proteins in-
dicates that the optimization of C4 function is much more 
complicated and may depend on lineage-specific physiological de-
tails. During the evolution of C4 biochemistry, the specific com-
partmentalization of enzyme activities and increased protein 
abundances created a new cellular environment (Lyu et al. 
2021). In addition, the evolution of different biochemical and ana-
tomical C4 subtypes imposed different physiological constraints 
between the C4 lineages. As a result, C4 enzymes coordinately 
adapted to the new metabolic context and the need for high car-
bon flux by fine-tuning their function through lineage-specific 
amino acid substitutions. An illustration of this process was pro-
vided by Lyu et al. (2021), who found that several genes encoding 
proteins associated with the C4 pathway in Flaveria species 
showed highly coordinated patterns of gene expression and mod-
ification in protein sequence.
The details of the molecular evolution of most C4 enzymes are 
still largely unknown. Present knowledge is based on the analysis 
of C4 enzymes from a small number of species in a few C4 lineages. 
Thus, it is not yet clear whether critical C4 changes are generally 
convergent, or whether they depend on the co-opted gene or on 
specific features of C4 subtypes. So, what adaptive amino acid sub-
stitutions make a protein suitable for C4 biochemistry in each lin-
eage? To answer this question, much more data from more 
species is needed to allow intra- and inter-lineage analyses to as-
sociate critical C4 changes with specific amino acid replacements.
Data on the kinetic and regulatory properties of an enzyme are 
rarely collected under conditions that mimic physiological ones, 
and they are therefore not sufficient to fully understand an 
Figure 11. Aspects of the C4 pathway. On the right: a schematic representation of the C4 biochemical pathway as found in maize and Flaveria. The main 
fluxes are indicated by thicker arrows. CBC, Calvin–Benson–Bassham Cycle. In the middle: 3D structural representations of tetrameric C4-PEPC (bottom) 
and C4-NADP-ME (top). On the left: subunits of C4-PEPC and C4-NADP-ME, highlighting differentially substituted amino acids. Amino acids in orange 
represent critical substitutions. Amino acids in black represent lineage-specific substitutions. The protein structures for Sorghum bicolor C4-NADP-ME 
(PDB code 6C7N) and Flaveria trinervia C4-PEPC (PDB code 3ZGE) were modeled using PyMOL Molecular Graphics System, version 2.0 Schrödinger, LLC.
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enzyme’s contribution to fitness. In fact, the performance of an 
enzyme depends on the physiological environment, emphasizing 
the importance of studying how adaptive changes affect plant 
physiology. At present, our knowledge of the in planta effects of 
adaptive changes is at a nascent stage. For example, in maize, 
Amaranthus, and Flaveria, the high activity level and low malate 
sensitivity of C4-PEPC were attributed to the phosphorylation state 
of the enzyme (Jiao and Chollet 1988; Tsuchida et al. 2001; Avasthi 
et al. 2011). However, the lack of phosphorylation of C4-PEPC from 
Flaveria bidentis was not crucial for PEPC activity or high photosyn-
thesis rates under standard greenhouse conditions (Furumoto 
et al. 2007). Another example is the pH-dependent inhibition by 
malate of C4-NADP-ME from maize, sorghum (Sorghum bicolor), 
and Setaria italica (Alvarez et al. 2019). The question is, how does 
this regulatory property—exclusive to the C4 isoform—affect 
plant physiology in a day-night cycle?
Several genetic engineering strategies to introduce a “C4-like 
pathway” into C3 plants have failed to improve photosynthesis, 
most likely due to the lack of adaptation of the C4 enzyme to the 
intracellular C3 environment. There is a need to assess the bio-
chemical and molecular characteristics of C4 enzymes in planta 
to understand how and under what conditions specific adaptive 
changes may affect plant physiology. In the future, new technol-
ogies such as CRISPR will facilitate the introduction of wild-type 
and mutated versions of C4 enzymes in C3 and C4 plants and the 
further analysis of their impact on plant metabolism and physiol-
ogy. For example, to assess whether lineage-specific adaptive 
changes are related to the optimization of C4 function, C4 mutant 
plants lacking their own photosynthetic C4-enzyme but express-
ing the C4 isoform of a distant lineage could be generated (e.g. 
C4-PEPC from grasses expressed in the C4 Flaveria trinervia 
C4-pepc
− mutant). In addition, C4 mutant plants expressing a dis-
tant C4 enzyme that contains amino acid substitutions specific 
to the lineage in which it is expressed could also be tested. 
These analyses would contribute to our overall understanding of 
the regulatory processes that are involved in the intricate dynam-
ics of C4 plant metabolism.
Conclusions
The questions covered here span a wide range of processes related 
to oxygenic photosynthesis. A few common themes and conclu-
sions can nonetheless be drawn from these diverse topics. First, 
protein interactions are critical and govern almost all cellular proc-
esses. The most exciting and important answers to unsolved ques-
tions will come from identifying the key proteins and their 
interacting partners, where and when the interactions take place, 
how they interact, and how they are regulated. A key aspect of 
many of the topics covered here is an emphasis on the importance 
of timing; when are molecular mechanisms active or interacting to-
gether? This is likely true for any biological system; therefore, an-
swers to these questions will provide fundamental insights not 
only into photosynthesis but many other biochemical processes 
as well. Second, the core photosystems PSI and PSII are highly con-
served among photosynthetic organisms, but many aspects of their 
regulation, from light harvesting to how the resulting chemical en-
ergy is dispersed and used in the cell and how excess energy is dis-
sipated, are highly diversified across the plant kingdom. Therefore, 
more work is needed not only on the historical model systems (e.g. 
Chlamydomonas, Arabidopsis) but on developing new model sys-
tems and understanding the diversity of photosynthesis in all its 
forms. Our discussions of the above questions further suggest 
that we are entering an era of understanding—and being able to 
interrogate—the ecophysiology of photosynthesis at the molecular 
level, which will be critical to understanding the diversity of photo-
synthetic regulatory mechanisms.
Acknowledgments
The authors apologize to those whose contributions could not be 
cited due to editorial restrictions. The authors thank Patrice 
Salomé for his excellent suggestions and work to improve figures, 
especially Figs. 5, 6, 7, 9, and 10. J.Y., V.K.Y., and J.K. would like to 
thank all the students, postdoctoral fellows, and collaborators for 
their contributions to their research. A.R.G. thanks Adrien 
Burlacot for useful recommendations on the alternative electron 
flow section. Y.A., H.M., and L.N. thank Dr. Dmitry Shevela 
(ShevelaDesign AB, Sweden) for the graphical work. M.H. thanks 
past and present Metaboactions team members for their hard 
work and stimulating discussions throughout the years. C.E.A., 
V.G.M., and M.A.T. thank Daria Chrobok (DC SciArt, Sweden) for 
professional graphic work for Fig. 11.
Author contributions
All authors contributed to writing and revising the article.
Funding
Work in the lab of J.Y., V.K.Y., and J.K. was supported by the U.S. 
Department of Energy, Office of Science,  Basic Energy Sciences), 
Chemical Sciences, Geosciences, and Biosciences Division 
(CSGB) (J.Y., V.K.Y., J.K.) for X-ray spectroscopy and crystallogra-
phy data collection and analysis and methods development for 
photosynthetic systems, by the National Institutes of Health) 
Grants 1R35GM149528 (V.K.Y.) for PS II biochemical studies, 
GM110501 (J.Y.) and GM126289 (J.K.) for instrumentation develop-
ment for XFEL experiments. Work by G.S.S.-C. and D.H. was 
supported by the U.S. Department of Energy, Office of Science, 
Basic Energy Sciences, Chemical Sciences, Geosciences, and 
Biosciences Division under Award #DE-SC0018097 to G.S.S.-C. 
Work by N.H. and C.B. was funded by the Deutsche 
Forschungsgemeinschaft (DFG, Bu812/8, Bu812/10) and supported 
by  Horizon 2020 of the European Union Research and Innovation 
Program ITN (grant no. 675006). The financial support of a UK 
Research and Innovation Future Leaders Fellowship (MR/ 
T017546/1 and MR/Y011635/1 to T.D.K. and T.C.) and an EPSRC 
Doctoral Training Partnership Scholarship (to E.C.) is gratefully 
acknowledged. Relevant work in the C.W.M. lab is supported 
by BBSRC (BB/W001012/1). Relevant work by W.W. and B.D.E. 
was supported by a Human Frontier Science Program Grant 
(RGP0005/2021) and ERC consolidator grant “cryOcean” (fulfilled 
by the Swiss State Secretariat for Education, Research and 
Innovation, M822.00045). Much of the work performed in the lab 
of A.R.G. on the export of metabolites from the chloroplast and al-
ternative electron transport pathways was supported by U.S. 
Department of Energy award DE-SC0019417.
Y.A. acknowledges Academy of Finland (project no. 315119). 
A.B. acknowledges the support of the Carnegie Institution 
for Science. Work in the team of M.H. has been supported by 
the Agence Nationale de la Recherche (ANR) (ANR-10- 
LABX-0040-SPS, ANR-11-IDEX-0003-02, ANR-14-CE19-0015 
REGUL3P), the CNRS, INRAE, University Paris-Saclay and the 
Saclay Plant Sciences Graduate School of Research. V.G.M. was 
funded by the Deutsche Forschungsgemeinschaft (DFG grant 
Open questions in photosynthesis research | 3933
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ctober 2024
MA2379/20-1 project number 441941117), and C.E.A. was funded 
by the ANPCyT grant PICT-2019-00079.
Conflict of interest statement. None declared.
References
Abadie C, Tcherkez G. Plant sulfur metabolism is stimulated by pho-
torespiration. Comm Biol. 2019:2(1):379. https://doi.org/10.1038/ 
s42003-019-0616-y
Alboresi A, Storti M, Cendron L, Morosinotto T. Role and regulation of 
class-C flavodiiron proteins in photosynthetic organisms. Biochem 
J. 2019a:476(17):2487–2498. https://doi.org/10.1042/BCJ20180648
Alboresi A, Storti M, Morosinotto T. Balancing protection and effi-
ciency in the regulation of photosynthetic electron transport 
across plant evolution. New Phytol. 2019b:221(1):105–109. 
https://doi.org/10.1111/nph.15372
Allahverdiyeva Y, Isojärvi J, Zhang P, Aro EM. Cyanobacterial oxy-
genic photosynthesis is protected by flavodiiron proteins. Life 
(Basel). 2015a:5(1):716–743. https://doi.org/10.3390/life5010716
Allahverdiyeva Y, Mustila H, Ermakova M, Bersanini L, Richaud P, 
Ajlani G, Battchikova N, Cournac L, Aro EM. Flavodiiron proteins 
Flv1 and Flv3 enable cyanobacterial growth and photosynthesis 
under fluctuating light. Proc. Natl. Acad. Sci. U S A. 2013:110(10): 
4111–4116. https://doi.org/10.1073/pnas.1221194110
Allahverdiyeva Y, Suorsa M, Tikkanen M, Aro EM. Photoprotection of 
photosystems in fluctuating light intensities. J Exp Bot. 2015b: 
66(9):2427–2436. https://doi.org/10.1093/jxb/eru463
Allen JF, Forsberg J. Molecular recognition in thylakoid structure and 
function. Trends Plant Sci. 2001:6(7):317–326. https://doi.org/10. 
1016/S1360-1385(01)02010-6
Allorent G, Tokutsu R, Roach T, Peers G, Cardol P, Girard-Bascou J, 
Seigneurin-Berny D, Petroutsos D, Kuntz M, Breyton C, et al. A 
dual strategy to cope with high light in Chlamydomonas reinhardtii. 
Plant Cell. 2013:25(2):545–557. https://doi.org/10.1105/tpc.112. 
108274
Alvarez CE, Bovdilova A, Hoppner A, Wolff CC, Saigo M, Trajtenberg 
F, Zhang T, Buschiazzo A, Nagel-Steger L, Drincovich MF, et al. 
Molecular adaptations of NADP-malic enzyme for its function 
in C4 photosynthesis in grasses. Nat Plants. 2019:5(7):755–765. 
https://doi.org/10.1038/s41477-019-0451-7
Alvarez CE, Maurino VG. Adaptive diversity in structure and function 
of C4 photosynthetic components. Biochem Soc Trans. 2023:51(3): 
1157–1168. https://doi.org/10.1042/BST20221279
Andersson B, Anderson JM. Lateral heterogeneity in the distribution 
of chlorophyll-protein complexes of the thylakoid membranes of 
spinach chloroplasts. Biochim Biophys Acta. 1980:593(2):427–440. 
https://doi.org/10.1016/0005-2728(80)90078-X
Appel J, Hueren V, Boehm M, Gutekunst K. Cyanobacterial in vivo so-
lar hydrogen production using a photosystem I–hydrogenase 
(PsaD-HoxYH) fusion complex. Nature Energy. 2020:5(6):458–467. 
https://doi.org/10.1038/s41560-020-0609-6
Armbruster U, Leonelli L, Correa Galvis V, Strand D, Quinn EH, 
Jonikas MC, Niyogi KK. Regulation and levels of the thylakoid K 
+/H+ antiporter KEA3 shape the dynamic response of photosyn-
thesis in fluctuating light. Plant Cell Physiol. 2016:57(7): 
1557–1567. https://doi.org/10.1093/pcp/pcw085
Arshad R, Calvaruso C, Boekema E, Büchel C, Kouřil R. Revealing the 
architecture of the photosynthetic apparatus in the diatom 
Thalassiosira pseudonana. Plant Physiol. 2021:186(4):2124–2136. 
https://doi.org/10.1093/plphys/kiab208
Asada K. THE WATER-WATER CYCLE IN CHLOROPLASTS: scaveng-
ing of active oxygens and dissipation of excess photons. Annu 
Rev Plant Physiol Plant Mol Biol. 1999:50(1):601–639. https://doi. 
org/10.1146/annurev.arplant.50.1.601
Avasthi UK, Izui K, Raghavendra AS. Interplay of light and tempera-
ture during the in planta modulation of C4 phosphoenolpyruvate 
carboxylase from the leaves of Amaranthus hypochondriacus L.: di-
urnal and seasonal effects manifested at molecular levels. J Exp 
Bot. 2011:62(3):1017–1026. https://doi.org/10.1093/jxb/erq333
Bailleul B, Rogato A, De Martino A, Coesel S, Cardol P, Bowler C, 
Falciatore A, Finazzi G. An atypical member of the light- 
harvesting complex stress-related protein family modulates dia-
tom responses to light. Proc Natl Acad Sci U S A. 2010:107(42): 
18214–18219. https://doi.org/10.1073/pnas.1007703107
Barber J, Andersson B. Too much of a good thing: light can be bad for 
photosynthesis. Trends Biochem Sci. 1992:17(2):61–66. https://doi. 
org/10.1016/0968-0004(92)90503-2
Barros T, Kühlbrandt W. Crystallisation, structure and function of 
plant light-harvesting Complex II. Biochim Biophys Acta. 2009: 
1787(6):753–772. https://doi.org/10.1016/j.bbabio.2009.03.012
Batool S, Uslu VV, Rajab H, Ahmad N, Waadt R, Geiger D, Malagoli M, 
Xiang CB, Hedrich R, Rennenberg H, et al. Sulfate is incorporated 
into cysteine to trigger ABA production and stomata closure. Plant 
Cell. 2018:30(12):2973–2987. https://doi.org/10.1105/tpc.18.00612
Bersanini L, Allahverdiyeva Y, Battchikova N, Heinz S, Lespinasse M, 
Ruohisto E, Mustila H, Nickelsen J, Vass I, Aro EM. Dissecting the 
photoprotective mechanism encoded by the flv4–2 operon: a dis-
tinct contribution of Sll0218 in photosystem II stabilization. Plant 
Cell Environ. 2017:40(3):378–389. https://doi.org/10.1111/pce.12872
Bersanini L, Battchikova N, Jokel M, Rehman A, Vass I, 
Allahverdiyeva Y, Aro EM. Flavodiiron protein Flv2/Flv4-related 
photoprotective mechanism dissipates excitation pressure of 
PSII in cooperation with phycobilisomes in Cyanobacteria. Plant 
Physiol. 2014:164(2):805–818. https://doi.org/10.1104/pp.113.231969
Bhattacharya D, Medlin L. The phylogeny of plastids: a review based 
on comparisons of small-subunit ribosomal RNA coding regions. J 
Phycol. 1995:31(4):489–498. https://doi.org/10.1111/j.1529-8817. 
1995.tb02542.x
Bhowmick A, Hussein R, Bogacz I, Simon PS, Ibrahim M, Chatterjee R, 
Doyle MD, Cheah MH, Fransson T, Chernev P, et al. Structural evi-
dence for intermediates during O2 formation in photosystem II. 
Nature. 2023:617(7961):629–636. https://doi.org/10.1038/s41586- 
023-06038-z
Bockwoldt M, Heiland I, Fischer K. The evolution of the plastid phos-
phate translocator family. Planta. 2019:250(1):245–261. https:// 
doi.org/10.1007/s00425-019-03161-y
Brown KA, Guo Z, Tokmina-Lukaszewska M, Scott LW, Lubner CE, 
Smolinski S, Mulder DW, Bothner B, King PW. The oxygen reduc-
tion reaction catalyzed by Synechocystis sp. PCC 6803 flavodiiron 
proteins. Sustain Energ. Fuels. 2019:3(11):3191–3200. https://doi. 
org/10.1039/c9se00523d
Büchel C. Evolution and function of light harvesting proteins. J. Plant 
Physiol. 2015:172(19):62–75. https://doi.org/10.1016/j.jplph.2014. 
04.018
Buchert F, Hamon M, Gabelein P, Scholz M, Hippler M, Wollman FA. 
The labile interactions of cyclic electron flow effector proteins. 
J Biol Chem. 2018:93(45):17559–17573. https://doi.org/10.1074/jbc. 
RA118.004475
Buck JM, Kroth PG, Lepetit B. Identification of sequence motifs in 
Lhcx proteins that confer qE-based photoprotection in the dia-
tom Phaeodactylum tricornutum. Plant J. 2021:108(6):1721–1734. 
https://doi.org/10.1111/tpj.15539
Burlacot A, Richaud P, Gosset A, Li-Beisson Y, Peltier G. Algal photo-
synthesis converts nitric oxide into nitrous oxide. Proc Natl Acad 
3934 | The Plant Cell, 2024, Vol. 36, No. 10
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Sci U S A. 2020:117(5):2704–2709. https://doi.org/10.1073/pnas. 
1915276117
Butkus V, Gelzinis A, Augulis R, Gall A, Büchel C, Robert B, Zigmantas 
D, Valkunas L, Abramavicius D. Coherence and population dy-
namics of chlorophyll excitations in FCP complex: two- 
dimensional spectroscopy study. J Chem Phys. 2015:142(21): 
212414. https://doi.org/10.1063/1.4914098
Calvaruso C, Rokka A, Aro E-M, Büchel C. Specific Lhc proteins are 
bound to PSI or PSII supercomplexes in the diatom Thalassiosira 
pseudonana. Plant Physiol. 2020:183(1):67–79. https://doi.org/10. 
1104/pp.20.00042
Cantrell M, Peers G. A mutant of Chlamydomonas without LHCSR 
maintains high rates of photosynthesis, but has reduced cell divi-
sion rates in sinusoidal light conditions. PLoS One. 2017:12(6): 
e0179395. https://doi.org/10.1371/journal.pone.0179395
Cardol P, Alric J, Girard-Bascou J, Franck F, Wollman FA, Finazzi G. 
Impaired respiration discloses the physiological significance of 
state transitions in Chlamydomonas. Proc Natl Acad Sci U S A. 
2009:106(37):15979–15984. https://doi.org/10.1073/pnas.0908111106
Carmo-Silva E, Sharwood RE. Rubisco and its regulation—major ad-
vances to improve carbon assimilation and productivity. J Exp Bot. 
2023:74(2):507–509. https://doi.org/10.1093/jxb/erac475
Casella S, Huang F, Mason D, Zhao G-Y, Johnson GN, Mullineaux CW, 
Liu L-N. Dissecting the native architecture and dynamics of cya-
nobacterial photosynthetic machinery. Mol Plant. 2017:10(11): 
1434–1448. https://doi.org/10.1016/j.molp.2017.09.019
Charuvi D, Nevo R, Kaplan-Ashiri I, Shimoni E, Reich Z. Studying the 
supramolecular organization of photosynthetic membranes 
within freeze-fractured leaf tissues by cryo-scanning electron 
microscopy. J Vis Exp. 2016(112):e54066. https://doi.org/10.3791/ 
54066
Charuvi D, Nevo R, Shimoni E, Naveh L, Zia A, Adam Z, Farrant JM, 
Kirchhoff H, Reich Z. Photoprotection conferred by changes in 
photosynthetic protein levels and organization during dehydra-
tion of a homoiochlorophyllous resurrection plant. Plant Physiol. 
2015:167(4):1554–1565. https://doi.org/10.1104/pp.114.255794
Chaux F, Burlacot A, Mekhalfi M, Auroy P, Blangy S, Richaud P, 
Peltier G. Flavodiiron proteins promote fast and transient O2 
photoreduction in Chlamydomonas. Plant Physiol. 2017:174(3): 
1825–1836. https://doi.org/10.1104/pp.17.00421
Chernev P, Fischer S, Hoffmann J, Oliver N, Assunção R, Yu B, Burnap 
RL, Zaharieva I, Nürnberg DJ, Haumann M, et al. Light-driven for-
mation of manganese oxide by today’s photosystem II supports 
evolutionarily ancient manganese-oxidizing photosynthesis. 
Nat Commun. 2020:11(1):6110. https://doi.org/10.1038/s41467- 
020-19852-0
Cho C, Ishii R, Hyeon S-B, Suzuki A. Stomatal regulation by aminoa-
cetonitrile, a photorespiration inhibitor. Plant Cell Physiol. 1987:28: 
1407–1410. https://doi.org/10.1093/oxfordjournals.pcp.a077432
Christin PA, Boxall SF, Gregory R, Edwards EJ, Hartwell J, Osborne CP. 
Parallel recruitment of multiple genes into c4 photosynthesis. 
Genome Biol. Evol. 2013:5(11):2174–2187. https://doi.org/10.1093/ 
gbe/evt168
Christin PA, Osborne CP. The evolutionary ecology of C4 plants. New 
Phytol. 2014:204(4):765–781. https://doi.org/10.1111/nph.13033
Christin PA, Salamin N, Savolainen V, Duvall MR, Besnard G. C4 pho-
tosynthesis evolved in grasses via parallel adaptive genetic 
changes. Curr Biol. 2007:17(14):1241–1247. https://doi.org/10. 
1016/j.cub.2007.06.036
Clausen CH, Brooks MD, Li T-D, Grob P, Kemalyan G, Nogales E, 
Niyogi KK, Fletcher DA. Dynamic mechanical responses of 
Arabidopsis thylakoid membranes during PSII-specific 
illumination. Biophys J. 2014:106(9):1864–1870. https://doi.org/10. 
1016/j.bpj.2014.03.016
Copley SD. Evolution of new enzymes by gene duplication and diver-
gence. FEBS J. 2020:287(7):1262–1283. https://doi.org/10.1111/febs. 
15299
Croce R, Carmo-Silva E, Cho YB, Ermakova M, Harbinson J, Lawson T, 
McCormick AJ, Niyogi KK, Ort DR, Patel-Tupper D. Perspectives on 
improving photosynthesis to increase crop yield. Plant Cell. 
2024:36(10):3944–3973. https://doi.org/10.1093/plcell/koae132
Croce R, van Amerongen H. Light harvesting in oxygenic photosyn-
thesis: structural biology meets spectroscopy. Science. 2020:369-
(6506):eaay2058. https://doi.org/10.1126/science.aay2058
Cruz JA, Savage LJ, Zegarac R, Hall CC, Satoh-Cruz M, Davis GA, 
Kovac WK, Chen J, Kramer DM. Dynamic environmental photo-
synthetic imaging reveals emergent phenotypes. Cell Syst. 
2016:2(6):365–377. https://doi.org/10.1016/j.cels.2016.06.001
Dai L, Tan LM, Jiang YL, Shi Y, Wang P, Zhang JP, Otomo ZY. 
Orientation assignment of LH2 and LH1-RC complexes from 
Thermochromatium tepidum reconstituted in PC liposome and 
their ultrafast excitation dynamics comparison between in artifi-
cial and in natural chromatophores. Chem Phys Lett. 2018:705: 
78–84. https://doi.org/10.1016/j.cplett.2018.05.043
Dalcorso G, Pesaresi P, Masiero S, Aseeva E, Schunemann D, Finazzi 
G, Joliot P, Barbato R, Leister D. A complex containing PGRL1 and 
PGR5 is involved in the switch between linear and cyclic electron 
flow in Arabidopsis. Cell. 2008:132(2):273–285. https://doi.org/10. 
1016/j.cell.2007.12.028
Dang KV, Plet J, Tolleter D, Jokel M, Cuine S, Carrier P, Auroy P, 
Richaud P, Johnson X, Alric J, et al. Combined increases in mito-
chondrial cooperation and oxygen photoreduction compensate 
for deficiency in cyclic electron flow in Chlamydomonas reinhardtii. 
Plant Cell. 2014:26(7):3036–3050. https://doi.org/10.1105/tpc.114. 
126375
Dao O, Kuhnert F, Weber APM, Peltier G, Li-Beisson Y. Physiological 
functions of malate shuttles in plants and algae. Trends Plant 
Sci. 2022:27(5):488–501. https://doi.org/10.1016/j.tplants.2021. 
11.007
Dau H, Haumann M. The manganese complex of photosystem II in 
its reaction cycle—basic framework and possible realization at 
the atomic level. Coord Chem Rev. 2008:252(3–4):273–295. https:// 
doi.org/10.1016/j.ccr.2007.09.001
Daum B, Nicastro D, Austin J 2nd, McIntosh JR, Kühlbrandt W. 
Arrangement of photosystem II and ATP synthase in chloroplast 
membranes of spinach and pea. Plant Cell. 2010:22(4):1299–1312. 
https://doi.org/10.1105/tpc.109.071431
Debus RJ. FTIR studies of metal ligands, networks of hydrogen bonds, 
and water molecules near the active site Mn4CaO5 cluster in pho-
tosystem II. Biochimi Biophys Acta. 2015:1847(1):19–34. https://doi. 
org/10.1016/j.bbabio.2014.07.007
Dekker JP, Boekema EJ. Supramolecular organization of thylakoid 
membrane proteins in green plants. Biochim Biophys Acta. 2005: 
1706(1–2):12–39. https://doi.org/10.1016/j.bbabio.2004.09.009
Dellero Y, Jossier M, Glab N, Oury C, Tcherkez G, Hodges M. 
Decreased glycolate oxidase activity leads to altered carbon allo-
cation and leaf senescence after a transfer from high CO2 to am-
bient air in Arabidopsis thaliana. J Exp Bot. 2016:67(10):3149–3163. 
https://doi.org/10.1093/jxb/erw054
Dellero Y, Lamothe-Sibold M, Jossier M, Hodges M. Arabidopsis thali-
ana ggt1 photorespiratory mutants maintain leaf carbon/nitrogen 
balance by reducing RuBisCO content and plant growth. Plant J. 
2015:83(6):1005–1118. https://doi.org/10.1111/tpj.12945
Depège N, Bellafiore S, Rochaix J-D. Role of chloroplast protein kinase 
stt7 in LHCII phosphorylation and state transition in 
Open questions in photosynthesis research | 3935
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Chlamydomonas. Science. 2003:299(5612):1572–1575. https://doi. 
org/10.1126/science.1081397
De Souza AP, Burgess SJ, Doran L, Hansen J, Manukyan L, Maryn N, 
Gotarkar D, Leonelli L, Niyogi KK, Long SP. Soybean photosynthe-
sis and crop yield are improved by accelerating recovery from 
photoprotection. Science. 2022:377(6608):851–854. https://doi.org/ 
10.1126/science.adc9831
Desplats C, Mus F, Cuine S, Billon E, Cournac L, Peltier G. 
Characterization of nda2, a plastoquinone-reducing type II 
NAD(P)H dehydrogenase in Chlamydomonas chloroplasts. J Biol 
Chem. 2009:284(7):4148–4157. https://doi.org/10.1074/jbc.M80454 
6200
Dietzel L, Bräutigam K, Pfannschmidt T. Photosynthetic acclimation: 
state transitions and adjustment of photosystem stoichiometry 
—functional relationships between short-term and long-term 
light quality acclimation in plants. FEBS J. 2008:275(6): 
1080–1088. https://doi.org/10.1111/j.1742-4658.2008.06264.x
Drincovich MF, Voll LM, Maurino VG. Editorial: on the diversity of 
roles of organic acids. Front Plant Sci. 2016:7:1592. https://doi. 
org/10.3389/fpls.2016.01592
Duminil P. 2019. Characterization of two primary metabolism en-
zymes in Arabidopsis thaliana: phosphoglycerate mutase and 
phosphoglycolate phosphatase [PhD Dissertation] University 
Paris-Saclay.
Durnford DG, Price JA, McKim SM, Sarchfield ML. Light-harvesting 
complex gene expression is controlled by both transcriptional 
and post-transcriptional mechanisms during photoacclimation 
in Chlamydomonas reinhardtii. Phys Plantarum. 2003:118(2): 
193–205. https://doi.org/10.1034/j.1399-3054.2003.00078.x
Eberhard S, Finazzi G, Wollman FA. The dynamics of photosynthesis. 
Ann Rev Genet. 2008:42(1):463–515. https://doi.org/10.1146/ 
annurev.genet.42.110807.091452
Eicks M, Maurino V, Knappe S, Flugge UI, Fischer K. The plastidic pen-
tose phosphate translocator represents a link between the cyto-
solic and the plastidic pentose phosphate pathways in plants. 
Plant Physiol. 2002:128(2):512–522. https://doi.org/10.1104/pp. 
010576
Eisenhut M, Bräutigam A, Timm S, Florian A, Tohge T, Fernie AR, 
Bauwe H, Weber APM. Photorespiration is crucial for dynamic re-
sponse of photosynthetic metabolism and stomatal movement to 
altered CO2 availability. Mol Plant. 2017:10(1):47–61. https://doi. 
org/10.1016/j.molp.2016.09.011
Eisenhut M, Georg J, Klähn S, Sakurai I, Mustila H, Zhang P, Hess WR, 
Aro EM. The antisense RNA as1_flv4 in the cyanobacterium 
Synechocystis sp. PCC 6803 prevents premature expression of the 
flv4–2 operon upon shift in inorganic carbon supply. J Biol Chem. 
2012:287(40):33153–33162. https://doi.org/10.1074/jbc.M112.391755
Eisenhut M, Roell M, Weber APM. Mechanistic understanding of pho-
torespiration paves the way to a new green revolution. New Phytol. 
2019:223(4):1762–1769. https://doi.org/10.1111/nph.15872
Engel BD, Schaffer M, Kuhn Cuellar L, Villa E, Plitzko JM, Baumeister 
W. Native architecture of the Chlamydomonas chloroplast re-
vealed by in situ cryo-electron tomography. Elife. 2015:4:e04889. 
https://doi.org/10.7554/eLife.04889
Ermakova M, Battchikova N, Allahverdiyeva Y, Aro EM. Novel 
heterocyst-specific flavodiiron proteins in Anabaena sp. PCC 
7120. FEBS Lett. 2013:587(1):82–87. https://doi.org/10.1016/j. 
febslet.2012.11.006
Ermakova M, Battchikova N, Richaud P, Leino H, Kosourov S, Isojärvi 
J, Peltier G, Flores E, Cournac L, Allahverdiyeva Y, et al. 
Heterocyst-specific flavodiiron protein Flv3B enables oxic diazo-
trophic growth of the filamentous cyanobacterium Anabaena sp. 
PCC 7120. Proc Natl Acad Sci USA. 2014:111(30):11205–11210. 
https://doi.org/10.1073/pnas.1407327111
Faries KM, Kressel LL, Wander MJ, Holten D, Laible PD, Kirmaier C, 
Hanson DK. High throughput engineering to revitalize a vestigial 
electron transfer pathway in bacterial photosynthetic reaction 
centers. J Biol Chem. 2012:287(11):8507–8514. https://doi.org/10. 
1074/jbc.M111.326447
Feng L, Raza MA, Li Z, Chen Y, Khalid MHB, Du J, Liu W, Wu X, Song C, 
Yu L, et al. The influence of light intensity and leaf movement on 
photosynthesis characteristics and carbon balance of soybean. 
Front Plant Sci. 2019:9:1952. https://doi.org/10.3389/fpls.2018. 
01952
Feng Y, Li Z, Li X, Shen L, Liu X, Zhou C, Zhang J, Sang M, Han G, Yang 
W, et al. Structure of a diatom photosystem II supercomplex con-
taining a member of lhcx family and dimeric FCPII. Sci Adv. 
2023:9(43):eadi8446. https://doi.org/10.1126/sciadv.adi8446
Fiebig OC, Harris D, Wang D, Hoffmann M, Schlau-Cohen GS. 
Ultrafast dynamics of photosynthetic light harvesting: strategies 
for acclimation across organisms. Annu Rev Phys Chem. 2023:74(1): 
493–520. https://doi.org/10.1146/annurev-physchem-083122-11 
1318
Field CB, Behrenfeld MJ, Randerson JT, Falkowski P. Primary produc-
tion of the biosphere: integrating terrestial and oceanic compo-
nents. Science. 1998:281(5374):237–240. https://doi.org/10.1126/ 
science.281.5374.237
Fischer K, Kammerer B, Gutensohn M, Arbinger B, Weber A, Hausler 
RE, Flugge UI. A new class of plastidic phosphate translocators: a 
putative link between primary and secondary metabolism by the 
phosphoenolpyruvate/phosphate antiporter. Plant Cell. 1997:199: 
453–462. https://doi.org/10.1105/tpc.9.3.453
Fleming GR, Schlau-Cohen GS, Amarnath K, Zaks J. Design principles 
of photosynthetic light-harvesting. Faraday Discuss. 2012:155: 
27–41. https://doi.org/10.1039/C1FD00078K
Flügel F, Timm S, Arrivault S, Florian A, Stitt M, Fernie AR, Bauwe H. 
The photorespiratory metabolite 2-phosphoglycolate regulates 
photosynthesis and starch accumulation in Arabidopsis. Plant 
Cell. 2017:29(10):2537–2551. https://doi.org/10.1105/tpc.17.00256
Flügge U-I, Häusler RE, Ludewig F, Fischer K. Functional genomics of 
phosphate antiport systems of plastids. Physiol Plant. 2003:118(4): 
475–482. https://doi.org/10.1034/j.1399-3054.2003.00137.x
Foudree A, Putarjunan A, Kambakam S, Nolan T, Fussell J, Pogorelko 
G, Rodermel S. The mechanism of variegation in immutans pro-
vides insight into chloroplast biogenesis. Front Plant Sci. 2012:3: 
260. https://doi.org/10.3389/fpls.2012.00260
Fu X, Gregory LM, Weise SE, Walker BJ. Integrated flux and pool size 
analysis in plant central metabolism reveals unique roles of gly-
cine and serine during photorespiration. Nat Plants. 2023:9(1): 
169–178. https://doi.org/10.1038/s41477-022-01294-9
Furumoto T, Izui K, Quinn V, Furbank RT, von Caemmerer S. 
Phosphorylation of phosphoenolpyruvate carboxylase is not es-
sential for high photosynthetic rates in the C4 species Flaveria bi-
dentis. Plant Physiol. 2007:144(4):1936–1945. https://doi.org/10. 
1104/pp.107.102541
Gan F, Zhang S, Rockwell NC, Martin SS, Lagarias JC, Bryant DA. 
Extensive remodeling of a cyanobacterial photosynthetic appara-
tus in far-red light. Science. 2014:345(6202):1312–1317. https://doi. 
org/10.1126/science.1256963
Gehlen J, Panstruga R, Smets H, Merkelbach S, Kleines M, Porsch P, 
Fladung M, Becker I, Rademacher T, Häusler RE, et al. Effects of 
altered phosphoenolpyruvate carboxylase activities on transgen-
ic C3 plant Solanum tuberosum. Plant Mol Biol. 1996:32(5):831–848. 
https://doi.org/10.1007/BF00020481
3936 | The Plant Cell, 2024, Vol. 36, No. 10
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Gerotto C, Alboresi A, Meneghesso A, Jokel M, Suorsa M, Aro EM, 
Morosinotto T. Flavodiiron proteins act as safety valve for elec-
trons in Physcomitrella patens. Proc Natl Acad Sci U S A. 2016:113-
(43):12322–12327. https://doi.org/10.1073/pnas.1606685113
Ghazaryan A, Akhtar P, Garab G, Lambrev PH, Büchel C. Involvement 
of the Lhcx protein Fcp6 of the diatom Cyclotella meneghiniana in 
the macro-organization and structural flexibility of thylakoid 
membranes. BBA-Bioenergetics. 2016:1857(9):1373–1379. https:// 
doi.org/10.1016/j.bbabio.2016.04.288
Gisriel CJ, Bryant DA, Brudvig GW, Cardona T. Molecular diversity 
and evolution of far-red light-acclimated photosystem I. Front 
Plant Sci. 2023:14:1289199. https://doi.org/10.3389/fpls.2023. 
1289199
Glatzel P, Bergmann U, Yano J, Visser H, Robblee JH, Gu WW, de Groot 
FMF, Christou G, Pecoraro VL, Cramer SP, et al. The electronic 
structure of mn in oxides, coordination complexes, and the 
oxygen-evolving complex of photosystem II studied by resonant 
inelastic X-ray scattering. J Am Chem Soc. 2004:126(32): 
9946–9959. https://doi.org/10.1021/ja038579z
Glatzel P, Schroeder H, Pushkar Y, Boron T III, Mukherjee S, Christou 
G, Pecoraro VL, Messinger J, Yachandra VK, Bergmann U, et al. 
Electronic structural changes of mn in the oxygen-evolving 
Complex of photosystem II during the catalytic cycle. Inorg 
Chem. 2013:52(10):5642–5644. https://doi.org/10.1021/ic4005938
Goral TK, Johnson MP, Brain APR, Kirchhoff H, Ruban AV, Mullineaux 
CW. Visualizing the mobility and distribution of chlorophyll pro-
teins in higher plant thylakoid membranes: effects of photoinhi-
bition and protein phosphorylation. Plant J. 2010:62(6):948–959. 
https://doi.org/10.1111/j.0960-7412.2010.04207.x
Goral TK, Johnson M, Duffy CDP, Brain APR, Ruban AV, Mullineaux 
CW. Light-harvesting antenna composition controls the 
macrostructure and dynamics of thylakoid membranes in 
Arabidopsis. Plant J. 2012:69(2):289–301. https://doi.org/10.1111/j. 
1365-313X.2011.04790.x
Greife P, Schönborn M, Capone M, Assunção R, Narzi D, Guidoni L, 
Dau H. The electron–proton bottleneck of photosynthetic oxygen 
evolution. Nature. 2023:617(7961):623–628. https://doi.org/10. 
1038/s41586-023-06008-5
Grouneva I, Jakob T, Wilhelm C, Goss R. A new multicomponent NPQ 
mechanism in the diatom Cyclotella meneghiniana. Plant Cell Physiol. 
2008:49(8):1217–1225. https://doi.org/10.1093/pcp/pcn097
Gundermann K, Büchel C. Factors determining the fluorescence 
yield of fucoxanthin-chlorophyll complexes (FCP) involved in 
non-photochemical quenching in diatoms. BBA-Bioenergetics. 
2012:1817(7):1044–1052. https://doi.org/10.1016/j.bbabio.2012. 
03.008
Gundermann K, Schmidt M, Weisheit W, Mittag M, Büchel C. 
Identification of several sub-populations in the pool of light har-
vesting proteins in the pennate diatom Phaeodactylum tricornutum. 
BBA-Bioenergetics. 2013:1827(3):303–310. https://doi.org/10.1016/j. 
bbabio.2012.10.017
Guo J, Nguyen AY, Dai Z, Su D, Gaffrey MJ, Moore RJ, Jacobs JM, 
Monroe ME, Smith RD, Koppenaal DW, et al. Proteome-wide 
light/dark modulation of thiol oxidation in cyanobacteria re-
vealed by quantitative site-specific redox proteomics. Mol Cell 
Proteomics. 2014:13(12):3270–3285. https://doi.org/10.1074/mcp. 
M114.041160
Harris D, Toporik H, Schlau-Cohen GS, Mazor Y. Energetic robust-
ness to large scale structural fluctuations in a photosynthetic 
supercomplex. Nat Commun. 2023:14(1):4650. https://doi.org/10. 
1038/s41467-023-40146-8
Hausler RE, Schlieben NH, Schulz B, Flugge UI. Compensation of de-
creased triose phosphate/phosphate translocator activity by 
accelerated starch turnover and glucose transport in transgenic 
tobacco. Planta. 1998:204(3):366–376. https://doi.org/10.1007/ 
s004250050268
He C, Berkowitz O, Hu S, Zhao Y, Qian K, Shou H, Whelan J, Wang J. 
Co-regulation of mitochondrial and chloroplast function: molec-
ular components and mechanisms. Plant Comm. 2023:4(1):100496. 
https://doi.org/10.1016/j.xplc.2022.100496
He S, Chou HT, Matthies D, Wunder T, Meyer MT, Atkinson N, 
Martinez-Sanchez A, Jeffrey PD, Port SA, Patena W, et al. The 
structural basis of Rubisco phase separation in the pyrenoid. 
Nat Plants. 2020:6(12):1480–1490. https://doi.org/10.1038/s41477- 
020-00811-y
Helman Y, Tchernov D, Reinhold L, Shibata M, Ogawa T, Schwarz R, 
Ohad I, Kaplan A. Genes encoding A-type flavoproteins are essen-
tial for photoreduction of O2 in cyanobacteria. Curr Biol. 2003: 
13(3):230–235. https://doi.org/10.1016/s0960-9822(03)00046-0
Herdean A, Teardo E, Nilsson AK, Pfeil BE, Johansson ON, Ünnep R, 
Nagy G, Zsiros O, Dana S, Solymosi K, et al. A voltage-dependent 
chloride channel fine-tunes photosynthesis in plants. Nat 
Commun. 2016:7(1):11654. https://doi.org/10.1038/ncomms11654
Horrer D, Flütsch S, Pazmino D, Matthews JS, Thalmann M, Nigro A, 
Leonhardt N, Lawson T, Santelia D. Blue light induces a distinct 
starch degradation pathway in guard cells for stomatal opening. 
Curr Biol. 2016:26(3):362–370. https://doi.org/10.1016/j.cub.2015. 
12.036
Houille-Vernes L, Rappaport F, Wollman FA, Alric J, Johnson X. 
Plastid terminal oxidase 2 (PTOX2) is the major oxidase involved 
in chlororespiration in Chlamydomonas. Proc Natl Acad Sci U S A. 
2011:108(51):20820–20825. https://doi.org/10.1073/pnas.111051 
8109
Huang W, Krishnan A, Plett A, Meagher M, Linka N, Wang Y, Ren B, 
Findinier J, Redekop P, Fakhima N, et al. Chlamydomonas mu-
tants lacking chloroplast TRIOSE PHOSPHATE TRANSPORTER3 
are metabolically compromised and light sensitive. Plant Cell. 
2023:35(7):2592–2614. https://doi.org/10.1093/plcell/koad095
Hüdig M, Tronconi MA, Zubimendi JP, Sage TL, Poschmann G, Bickel 
D, Gohlke H, Maurino VG. Respiratory and C4-photosynthetic 
NAD-malic enzyme coexist in bundle sheath cell mitochondria 
and evolved via association of differentially adapted subunits. 
Plant Cell. 2022:34(1):597–615. https://doi.org/10.1093/plcell/ 
koab265
Huokko T, Ni T, Dykes GF, Simpson DM, Brownridge P, Conradi FD, 
Beynon RJ, Nixon PJ, Mullineaux CW, Zhang P, et al. Probing the 
biogenesis pathway and dynamics of thylakoid membranes. 
Nat Commun. 2021:12(1):3475. https://doi.org/10.1038/s41467- 
021-23680-1
Hussein R, Ibrahim M, Bhowmick A, Simon PS, Bogacz I, Doyle MD, 
Dobbek H, Zouni A, Messinger J, Yachandra VK, et al. 
Evolutionary diversity of proton and water channels on the oxi-
dizing side of photosystem II and their relevance to function. 
Photosynth. Res. 2023:158(2):91–107. https://doi.org/10.1007/ 
s11120-023-01018-w
Hussein R, Ibrahim M, Bhowmick A, Simon PS, Chatterjee R, Lassalle 
L, Doyle M, Bogacz I, Kim IS, Cheah MH, et al. Structural dynamics 
in the water and proton channels of photosystem II during the S2 
to S3 transition. Nat Commun. 2021:12(1):6531. https://doi.org/10. 
1038/s41467-021-26781-z
Ibrahim M, Fransson T, Chatterjee R, Cheah MH, Hussein R, Lassalle 
L, Sutherlin KD, Young ID, Fuller FD, Gul S, et al. Untangling the 
sequence of events during the S2 → S3 transition in photosystem 
II and implications for the water oxidation mechanism. Proc Natl 
Acad Sci U S A. 2020:117(23):12624–12635. https://doi.org/10.1073/ 
pnas.2000529117
Open questions in photosynthesis research | 3937
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Ilík P, Pavlovič A, Kouřil R, Alboresi A, Morosinotto T, Allahverdiyeva 
Y, Aro EM, Yamamoto H, Shikanai T. Alternative electron trans-
port mediated by flavodiiron proteins is operational in organisms 
from cyanobacteria up to gymnosperms. New Phytol. 2017:214(3): 
967–972. https://doi.org/10.1111/nph.14536
Ishikita H, Saenger W, Biesiadka J, Loll B, Knapp EW. How photosyn-
thetic reaction centers control oxidation power in chlorophyll 
pairs P680, P700, and P870. Proc Natl Acad Sci U S A. 2006:103(26): 
9855–9860. https://doi.org/10.1073/pnas.0601446103
Jackson PJ, Hitchcock A, Brindley AA, Dickman MJ, Hunter CN. 
Absolute quantification of cellular levels of photosynthesis- 
related proteins in Synechocystis sp. PCC 6803. Photosynth Res. 
2023:155(3):219–245. https://doi.org/10.1007/s11120-022-00990-z
Jiao JA, Chollet R. Light/dark regulation of maize leaf phosphoenol-
pyruvate carboxylase by in vivo phosphorylation. Arch Biochem 
Biophys. 1988:261(2):409–417. https://doi.org/10.1016/0003-9861 
(88)90357-8
Joet T, Cournac L, Horvath EM, Medgyesy P, Peltier G. Increased sen-
sitivity of photosynthesis to antimycin A induced by inactivation 
of the chloroplast ndhB gene. Evidence for a participation of the 
NADH-dehydrogenase complex to cyclic electron flow around 
photosystem I. Plant Physiol. 2001:125(4):1919–1929. https://doi. 
org/10.1104/pp.125.4.1919
Jokel M, Johnson X, Peltier G, Aro EM, Allahverdiyeva Y. Hunting the 
main player enabling Chlamydomonas reinhardtii growth under 
fluctuating light. Plant J. 2018:94(5):822–835. https://doi.org/10. 
1111/tpj.13897
Joliot P, Selles J, Wollman FA, Vermeglio A. High efficient cyclic elec-
tron flow and functional supercomplexes in Chlamydomonas 
cells. Biochim Biophys Acta Bioenerg. 2022:1863(8):148909. https:// 
doi.org/10.1016/j.bbabio.2022.148909
Kammerer B, Fischer K, Hilpert B, Scubert S, Gutensohn M, Weber A, 
Flugge UI. Molecular characterization of a carbon transporter in 
plastids from heterotrophic tissues: the glucose 6-phosphate/ 
phosphate antiporter. Plant Cell. 1998:10(1):105–117. https://doi. 
org/10.1105/tpc.10.1.105
Kanygin A, Milrad Y, Thummala C, Reifschneider K, Baker P, Marco P, 
Yacoby I, Redding KE. Rewiring photosynthesis: a photosystem 
I-hydrogenase chimera that makes H2 in vivo. Energy Environ 
Sci. 2020:13(9):2903–2914. https://doi.org/10.1039/C9EE03859K
Kaye Y, Huang W, Clowez S, Saeoussi S, Idoine A, Sanz-Luque E, 
Grossman AR. The mitochondrial alternative oxidase from 
Chlamydomonas reinhardtii enables survival in high light. J Biol 
Chem. 2019:294(4):1380–1395. https://doi.org/10.1074/jbc.RA118. 
004667
Kern J, Alonso-Mori R, Hellmich J, Rosalie T, Hattne J, Laksmono H, 
Gloeckner C, Echols N, Sierra RG, Sellberg J, et al. Room temper-
ature femtosecond X-ray diffraction of photosystem II microcrys-
tals. Proc Natl Acad Sci U S A. 2012:109(25):9721–9726. https://doi. 
org/10.1073/pnas.1204598109
Kern J, Alonso-Mori R, Tran R, Hattne J, Gildea RJ, Echols N, Glockner 
C, Hellmich J, Laksmono H, Sierra RG, et al. Simultaneous femto-
second X-ray spectroscopy and diffraction of photosystem II at 
room temperature. Science. 2013:340(6131):491–495. https://doi. 
org/10.1126/science.1234273
Kern J, Tran R, Alonso-Mori R, Koroidov S, Echols N, Hattne J, Ibrahim 
M, Gul S, Laksmono H, Sierra RG, et al. Taking snapshots of photo-
synthetic water oxidation using femtosecond X-ray diffraction 
and spectroscopy. Nature Commun. 2014:5(1):4371. https://doi. 
org/10.1038/ncomms5371
Keskin O, Tuncbag N, Gursoy A. Predicting protein–protein interac-
tions from the molecular to the proteome level. Chem Rev. 2016: 
116(8):4884–4909. https://doi.org/10.1021/acs.chemrev.5b00683
Kirchhoff H. Diffusion of molecules and macromolecules in thyla-
koid membranes. Biochim Biophys Acta. 2014a:1837(4):495–502. 
https://doi.org/10.1016/j.bbabio.2013.11.003
Kirchhoff H. Structural changes of the thylakoid membrane network 
induced by high light stress in plant chloroplasts. Philosophical 
Transactions of the Royal Society B: Biological Sciences. 
2014b:369(1640):20130225. https://doi.org/10.1098/rstb.2013. 
0225
Kirchhoff H, Haase W, Wegner S, Danielsson R, Ackermann R, 
Albertsson P-A. Low-light-induced formation of semicrystalline 
photosystem II arrays in higher plant chloroplasts. Biochem. 
2007:46(39):11169–11176. https://doi.org/10.1021/bi700748y
Kirchhoff H, Haferkamp S, Allen JF, Epstein DBA, Mullineaux CW. 
Protein diffusion and macromolecular crowding in thylakoid 
membranes. Plant Physiol. 2008:146(4):1571–1578. https://doi.org/ 
10.1104/pp.107.115170
Kirchhoff H, Mukherjee U, Galla HJ. Molecular architecture of the 
thylakoid membrane: lipid diffusion space for plastoquinone. 
Biochem. 2002:41(15):4872–4882. https://doi.org/10.1021/bi011650y
Kok B, Forbush B, McGloin M. Cooperation of charges in photosyn-
thetic oxygen evolution. A linear four step mechanism. 
Photochem Photobiol. 1970:11(6):457–475. https://doi.org/10.1111/j. 
1751-1097.1970.tb06017.x
Komenda J, Sobotka R, Nixon PJ. Assembling and maintaining the 
photosystem II complex in chloroplasts and cyanobacteria. Curr 
Opin Plant Biol. 2012:15(3):245–251. https://doi.org/10.1016/j.pbi. 
2012.01.017
Kromdijk J, Głowacka K, Leonelli L, Gabilly ST, Iwai M, Niyogi KK, 
Long SP. Improving photosynthesis and crop productivity by ac-
celerating recovery from photoprotection. Science. 2016:354-
(6314):857–861. https://doi.org/10.1126/science.aai8878
Kumazawa M, Nishide H, Nagao R, Inoue-Kashino N, Shen J-R, 
Nakano T, Uchiyama I, Kashino Y, Ifuku K. Molecular phylogeny 
of fucoxanthin-chlorophyll a/c proteins from Chaetoceros gracilis 
and Lhcq/Lhcf diversity. Physiol. Plantarum. 2022:174(1):e13598. 
https://doi.org/10.1111/ppl.13598
Lavaud J, Rousseau B, van Gorkom HJ, Etienne A-L. Influence of the 
diadinoxanthin pool size on photoprotection in the marine plank-
tonic diatom Phaeodactylum tricornutum. Plant Physiol. 2002:129(3): 
1398–1406. https://doi.org/10.1104/pp.002014
Lemaire C, Wollman FA, Bennoun P. Restoration of phototrophic 
growth in a mutant of Chlamydomonas reinhardtii in which the 
chloroplast atpB gene of the ATP synthase has a deletion: an ex-
ample of mitochondria-dependent photosynthesis. Proc Natl 
Acad Sci U S A. 1988:85(5):1344–1348. https://doi.org/10.1073/ 
pnas.85.5.1344
Lemonnier P, Lawson T. Calvin cycle and guard cell metabolism im-
pact stomatal function. Semin Cell Dev Biol. 2023:155(Pt A):59–70. 
https://doi.org/10.1016/j.semcdb.2023.03.001
Levitan O, Chen M, Kuang X, Cheong KY, Jiang J, Banal M, Nambiar N, 
Gorbunov MY, Ludtke SJ, Falkowski PG, et al. Structural and func-
tional analyses of photosystem II in the marine diatom 
Phaeodactylum tricornutum. Proc Natl Acad Sci U S A. 2019:116-
(35):17316–17322. https://doi.org/10.1073/pnas.1906726116
Li M, Ma J, Li X, Sui S-F. In situ cryo-ET structure of phycobilisome– 
photosystem II supercomplex from red alga. eLife. 2021a:10: 
e69635. https://doi.org/10.7554/eLife.69635
Li M, Svoboda V, Davis G, Kramer D, Kunz H-H, Kirchhoff H. Impact of 
ion fluxes across thylakoid membranes on photosynthetic elec-
tron transport and photoprotection. Nat Plants. 2021b:7(7): 
979–988. https://doi.org/10.1038/s41477-021-00947-5
Lim SL, Flütsch S, Liu J, Distefano L, Santelia D, Lim BL. Guard cell 
chloroplasts import cytosolic ATP for starch turnover and 
3938 | The Plant Cell, 2024, Vol. 36, No. 10
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
stomatal opening. Nat Commun. 2022:13(1):652. https://doi.org/10. 
1038/s41467-022-28263-2
Lindahl M, Spetea C, Hundal T, Oppenheim AB, Adam Z, Andersson 
B. The thylakoid FtsH protease plays a role in the light-induced 
turnover of the photosystem II D1 protein. Plant Cell. 2000:12(3): 
419–431. https://doi.org/10.1105/tpc.12.3.419
Liu Y, Mauve C, Lamothe-Sibold M, Guérard F, Glab N, Hodges M, 
Jossier M. Photorespiratory serine hydroxymethyltransferase 1 
activity impacts abiotic stress tolerance and stomatal closure. 
Plant Cell Environ. 2019:42(9):2567–2583. https://doi.org/10.1111/ 
pce.13595
López-Calcagno PE, Fisk S, Brown KL, Bull SE, South PF, Raines CA. 
Overexpressing the H-protein of the glycine cleavage system in-
creases biomass yield in glasshouse and field-grown transgenic 
tobacco plants. Plant Biotechnol J. 2019:17(1):141–151. https://doi. 
org/10.1111/pbi.12953
Lu Y, Li Y, Yang Q, Zhang Z, Chen Y, Zhang S, Peng XX. Suppression of 
glycolate oxidase causes glyoxylate accumulation that inhibits 
photosynthesis through deactivating rubisco in rice. Physiol 
Plant. 2014:150(3):463–476. https://doi.org/10.1111/ppl.12104
Lubitz W, Chrysina M, Cox N. Water oxidation in photosystem II. 
Photosynth. Res. 2019:142(1):105–125. https://doi.org/10.1007/ 
s11120-019-00648-3
Lüer L, Moulisová V, Henry S, Polli D, Brotosudarmo THP, 
Hoseinkhani S, Brida D, Lanzani G, Cerullo G, Cogdell RJ. 
Tracking energy transfer between light harvesting complex 2 
and 1 in photosynthetic membranes grown under high and low 
illumination. Proc Natl Acad Sci U S A. 2012:109(5):1473–1478. 
https://doi.org/10.1073/pnas.1113080109
Lyu MA, Gowik U, Kelly S, Covshoff S, Hibberd JM, Sage RF, Ludwig M, 
Wong GK, Westhoff P, Zhu XG. The coordination of major events 
in C(4) photosynthesis evolution in the genus Flaveria. Scientific 
Rep. 2021:11(1):15618. https://doi.org/10.1038/s41598-021-93381-8
MacGregor-Chatwin C, Nürnberg DJ, Jackson PJ, Vasilev C, Hitchcock 
A, Ho M-Y, Shen G, Gisriel CJ, Wood William HJ, Mahbub M. 
Changes in supramolecular organization of cyanobacterial thyla-
koid membrane complexes in response to far-red light photoac-
climation. Sci Adv. 2022:8(6). https://doi.org/10.1126/sciadv. 
abj4437
MacGregor-Chatwin C, Sener M, Barnett SFH, Hitchcock A, 
Barnhart-Dailey MC, Maghlaoui K, Barber J, Timlin JA, Schulten 
K, Hunter CN. Lateral segregation of photosystem I in cyanobac-
terial thylakoids. Plant Cell. 2017:29(5):1119–1136. https://doi.org/ 
10.1105/tpc.17.00071
Mahbub M, Hemm L, Yang Y, Kaur R, Carmen H, Engl C, Huokko T, 
Riediger M, Watanabe S, Liu L-N, et al. mRNA localization, reac-
tion centre biogenesis and thylakoid membrane targeting in cya-
nobacteria. Nat Plants. 2020:6(9):1179–1191. https://doi.org/10. 
1038/s41477-020-00764-2
Mahbub M, Mullineaux CW. Locations of membrane protein produc-
tion in a cyanobacterium. J Bacteriol. 2023:205(10):e00209–e00223. 
https://doi.org/10.1128/jb.00209-23
Maier A, Fahnenstich H, von Caemmerer S, Engqvist MKM, Weber 
APM, Flügge U-I, Maurino VG. Transgenic introduction of a glyco-
late oxidative cycle into A. thaliana chloroplasts leads to growth 
improvement. Front Plant Sci. 2012:3:38. https://doi.org/10.3389/ 
fpls.2012.00038
Maier A, Zell MB, Maurino VG. Malate decarboxylases: evolution and 
roles of NAD(P)-ME isoforms in species performing C(4) and C(3) 
photosynthesis. J Exp Bot. 2011:62(9):3061–3069. https://doi.org/ 
10.1093/jxb/err024
Makino A, Miyake C, Yokota A. Physiological functions of the water- 
water cycle (Mehler reaction) and the cyclic electron flow around 
PSI in rice leaves. Plant Cell Physiol. 2002:43(9):1017–1026. https:// 
doi.org/10.1093/pcp/pcf124
Malone LA, Proctor MS, Hitchcock A, Hunter CN, Johnson MP. 
Cytochrome b6f—orchestrator of photosynthetic electron trans-
fer. Biochim Biophys Acta. 2021:1862(5):148380. https://doi.org/10. 
1016/j.bbabio.2021.148380
Manna P, Hoffmann M, Davies T, Richardson KH, Johnson MP, 
Schlau-Cohen GS. Energetic driving force for LHCII clustering in 
plant membranes. Sci Adv. 2023:9(51):2023–2004. https://doi.org/ 
10.1126/sciadv.adj0807
Mao R, Zhang H, Bie L, Liu L-N, Gao J. Million-atom molecular dynam-
ics simulations reveal the interfacial interactions and assembly 
of plant PSII-LHCII supercomplex. RSC Adv. 2023:13(10): 
6699–6712. https://doi.org/10.1039/D2RA08240C
Mazor Y, Nataf D, Toporik H, Nelson N. Crystal structures of virus- 
like photosystem I complexes from the mesophilic cyanobacte-
rium Synechocystis PCC 6803. eLife. 2014:3:e01496. https://doi. 
org/10.7554/eLife.01496
Messant M, Timm S, Fantuzzi A, Weckwerth W, Bauwe H, Rutherford 
AW, Krieger-Liszkay A. Glycolate induces redox tuning of photo-
system II in vivo: study of a photorespiration mutant. Plant Physiol. 
2018:177(3):1277–1285. https://doi.org/10.1104/pp.18.00341
Mettler T, Mühlhaus T, Hemme D, Schöttler M-A, Rupprecht J, Idoine 
A, Veyel D, Pal SK, Yaneva-Roder L, Winck FV, et al. Systems anal-
ysis of the response of photosynthesis, metabolism, and growth 
to an increase in irradiance in the photosynthetic model organ-
ism Chlamydomonas reinhardtii. Plant Cell 2014:26(6): 2310–2350 
https://doi.org/10.1105/tpc.114.124537
Meyers BE, Boyd SP. Signal decomposition using masked proximal 
operators. Found Trend Signal Proc. 2023:17(1):1–78. https://doi. 
org/10.1561/2000000122
Mirkovic T, Ostroumov EE, Anna JM, van Grondelle R, Govindjee, 
Scholes GD. Light absorption and energy transfer in the antenna 
complexes of photosynthetic organisms. Chem Rev. 2017:117(2): 
249–293. https://doi.org/10.1021/acs.chemrev.6b00002
Morales A, Kaiser E. Photosynthetic acclimation to fluctuating irradi-
ance in plants. Front Plant Sci. 2020:11:268. https://doi.org/10.3389/ 
fpls.2020.00268
Munekage Y, Hashimoto M, Miyake C, Tomizawa K, Endo T, Tasaka 
M, Shikanai T. Cyclic electron flow around photosystem I is es-
sential for photosynthesis. Nature. 2004:429(6991):579–582. 
https://doi.org/10.1038/nature02598
Mustárdy L, Buttle K, Steinbach G, Garab G. The three-dimensional 
network of the thylakoid membranes in plants: quasihelical 
model of the granum-stroma assembly. Plant Cell. 2008:20(10): 
2552–2557. https://doi.org/10.1105/tpc.108.059147
Mustila H, Paananen P, Battchikova N, Santana-Sánchez A, 
Muth-Pawlak D, Hagemann M, Aro EM, Allahverdiyeva Y. The fla-
vodiiron protein Flv3 functions as a homo-oligomer during stress 
acclimation and is distinct from the Flv1/Flv3 hetero-oligomer 
specific to the O2 photoreduction pathway. Plant Cell Physiol. 
2016:57(7):1468–1483. https://doi.org/10.1093/pcp/pcw047
Nagao R, Kato K, Ifuku K, Suzuki T, Kumazawa M, Uchiyama I, 
Kashino Y, Dohmae N, Akimoto S, Shen J-R, et al. Structural basis 
for assembly and function of a diatom photosystem 
I-light-harvesting supercomplex. Nat Commun. 2020:11(1):2481. 
https://doi.org/10.1038/s41467-020-16324-3
Nagao R, Kitazaki S, Noguchi T. Evaluation of photosynthetic activ-
ities in thylakoid membranes by means of Fourier transform in-
frared spectroscopy. Biochim Biophys Acta Bioenerg. 2018:1859(2): 
129–136. https://doi.org/10.1016/j.bbabio.2017.11.004
Narusaka Y, Narusaka M, Kobayashi H, Satoh K. The herbicide- 
resistant species of the cyanobacterial D1 protein obtained by 
Open questions in photosynthesis research | 3939
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
thorough and random in vitro mutagenesis. Plant Cell Physiol. 
1998:39(6):620–626. https://doi.org/10.1093/oxfordjournals.pcp. 
a029413
Narusaka Y, Narusaka M, Satoh K, Kobayashi H. In vitro random mu-
tagenesis of the D1 protein of the photosystem II reaction center 
confers phototolerance on the cyanobacterium Synechocystis sp. 
PCC 6803*. J Biol Chem. 1999:274(33):23270–23275. https://doi.org/ 
10.1074/jbc.274.33.23270
Nawrocki WJ, Buchert F, Joliot P, Rappaport F, Bailleul B, Wollman 
F-A. Chlororespiration controls growth under intermittent light. 
Plant Physiol. 2019:179(2):630–639. 10.1104/pp.18.01213
Nayak L, Panda D, Dash GK, Lal MK, Swain P, Baig MJ, Kumar A. 
A chloroplast glycolate catabolic pathway bypassing the endoge-
nous photorespiratory cycle enhances photosynthesis, biomass 
and yield in rice (Oryza sativa L). Plant Sci. 2022:314:111103. 
https://doi.org/10.1016/j.plantsci.2021.111103
Nedbal L, Březina V. Complex metabolic oscillations in plants forced 
by harmonic irradiance. Biophys J. 2002:83(4):2180–2189. https:// 
doi.org/10.1016/S0006-3495(02)73978-7
Nedbal L, Lazár D. Photosynthesis dynamics and regulation sensed 
in the frequency domain. Plant Physiol. 2021:187(2):646–661. 
https://doi.org/10.1093/plphys/kiab317
Nelson N, Junge W. Structure and energy transfer in photosystems of 
oxygenic photosynthesis. Annu Rev Biochem. 2015:84(1):659–683. 
https://doi.org/10.1146/annurev-biochem-092914-041942
Nelson N, Yocum CF. Structure and function of photosystems I and 
II. Ann Rev Plant Biol. 2006:57(1):521–565. https://doi.org/10.1146/ 
annurev.arplant.57.032905.105350
Nevo R, Charuvi D, Shimoni E, Schwarz R, Kaplan A, Ohad I, Reich Z. 
Thylakoid membrane perforations and connectivity enable intra-
cellular traffic in cyanobacteria. EMBO J. 2007:26(5):1467–1473. 
https://doi.org/10.1038/sj.emboj.7601594
Nguyen TB, Lefoulon C, Nguyen TH, Blatt MR, Carroll W. Engineering 
stomata for enhanced carbon capture and water-use efficiency. 
Trends Plant Sci. 2023:28(11):1290–1309. https://doi.org/10.1016/j. 
tplants.2023.06.002
Nicol L, Croce R. The PsbS protein and low pH are necessary and suf-
ficient to induce quenching in the light-harvesting complex of 
plants LHCII. Sci Rep. 2021:11(1):7415. https://doi.org/10.1038/ 
s41598-021-86975-9
Nikkanen L, Santana Sánchez A, Ermakova M, Rögner M, Cournac L, 
Allahverdiyeva Y. Functional redundancy between flavodiiron 
proteins and NDH-1 in Synechocystis sp. PCC 6803. Plant J. 
2020:103(4):1460–1476. https://doi.org/10.1111/tpj.14812
Nikkanen L, Vakal S, Santana Sánchez A, Hubáček M, Wang Y, Böhm 
M, Gutekunst K, Salminen TA, Allahverdiyeva Y. Proton motive 
force dissipation drives flavodiiron proteins to the thylakoid 
membrane for ferredoxin-powered O2 photoreduction. bioRxiv 
541409, https://doi.org/10.1101/2023.05.19.541409, 22 May 2023, 
preprint: not peer reviewed.
Niu Y, Lazár D, Holzwarth AR, Kramer DM, Matsubara S, Fiorani F, 
Poorter H, Schrey SD, Nedbal L. Plants cope with fluctuating light 
by frequency-dependent nonphotochemical quenching and 
cyclic electron transport. New Phytol. 2023:239(5):1869–1886. 
https://doi.org/10.1111/nph.19083
Niyogi KK, Truong TB. Evolution of flexible non-photochemical 
quenching mechanisms that regulate light harvesting in oxy-
genic photosynthesis. Curr Opin Plant Biol. 2013:16(3):307–314. 
https://doi.org/10.1016/j.pbi.2013.03.011
Nunes-Nesi A, Carrari F, Gibon Y, Sulpice R, Lytovchenko A, Fisahn J, 
Graham J, Ratcliffe RG, Sweetlove LJ, Fernie AR. Deficiency of mi-
tochondrial fumarase activity in tomato plants impairs 
photosynthesis via an effect on stomatal function. Plant J. 
2007:50(6):1093–1106. https://doi.org/10.1111/j.1365-313X.2007. 
03115.x
Ojakian GK, Satir P. Particle movements in chloroplast membranes: 
quantitative measurements of membrane fluidity by the freeze- 
fracture technique. Proc Natl Acad Sci U S A. 1974:71(5): 
2052–2056. https://doi.org/10.1073/pnas.71.5.2052
Oliver T, Kim TD, Trinugroho JP, Cordón-Preciado V, Wijayatilake N, 
Bhatia A, Rutherford AW, Cardona T. The evolution and evolv-
ability of photosystem II. Annu Rev Plant Biol. 2023:74(1):225–257. 
https://doi.org/10.1146/annurev-arplant-070522-062509
Onoa B, Fukuda S, Iwai M, Bustamante C, Niyogi KK. Atomic force mi-
croscopy visualizes mobility of photosynthetic proteins in grana 
thylakoid membranes. Biophys J. 2020:118(8):1876–1886. https:// 
doi.org/10.1016/j.bpj.2020.02.029
Opatíková M, Semchonok DA, Kopečný D, Ilík P, Pospíšil P, Ilíková I, 
Roudnický P, Zeljković SC, Tarkowski P, Kyrilis FL, et al. 
Cryo-EM structure of a plant photosystem II supercomplex with 
light-harvesting protein Lhcb8 and α-tocopherol. Nat Plants. 
2023:9(8):1359–1369. https://doi.org/10.1038/s41477-023-01483-0
Papagiannakis E, van Stokkum IHM, Fey H, Büchel C, van Grondelle 
R. Spectroscopic characterization of the excitation energy trans-
fer in the fucoxanthin-chlorophyll protein of diatoms. Photosynth 
Res. 2005:86(1–2):241–250. https://doi.org/10.1007/s11120-005- 
1003-8
Park S, Steen CJ, Fischer AL, Fleming GR. Snapshot transient absorp-
tion spectroscopy: toward in vivo investigations of nonphoto-
chemical quenching mechanisms. Photosyn Res. 2019:141(3): 
367–376. https://doi.org/10.1007/s11120-019-00640-x
Pearcy RW. Sunflecks and photosynthesis in plant canopies. Annu 
Rev Plant Physiol Plant Mol Biol. 1990:41(1):421–453. https://doi. 
org/10.1146/annurev.pp.41.060190.002225
Peltier G, Aro EM, Shikanai T. NDH-1 and NDH-2 plastoquinone re-
ductases in oxygenic photosynthesis. Annu Rev Plant Biol. 2016: 
67(1):55–80. https://doi.org/10.1146/annurev-arplant-043014- 
114752
Peltier G, Stoffel C, Findinier J, Madireddi SK, Dao O, Epting V, Morin 
A, Grossman A, Li-Beisson Y, Burlacot A. Green algal CO2 capture 
is powered by alternative electron pathways of photosynthesis. 
bioRxiv 552514, https://doi.org/10.1101/2023.08.08.552514, 22 
November 2023, preprint: not peer reviewed.
Peltier G, Tolleter D, Billon E, Cournac L. Auxiliary electron transport 
pathways in chloroplasts of microalgae. Photosynth Res. 
2010:106(1–2):19–31. https://doi.org/10.1007/s11120-010-9575-3
Peng J, Lu J, Hoh D, Dina AS, Shang X, Kramer DM, Chen J. Identifying 
emerging phenomenon in long temporal phenotyping experi-
ments. Bioinformatics. 2020:36(2):568–577. https://doi.org/10. 
1093/bioinformatics/btz559
Perera-Castro AV, Flexas J. Recent advances in understanding and 
improving photosynthesis. Fac Rev. 2020:9:5. https://doi.org/10. 
12703/b/9-5
Perez-Boerema A, Engel BD, Wietrzynski W. Evolution of thylakoid 
structural diversity. Annu Rev Cell Dev Biol. https://doi.org/10. 
1146/annurev-cellbio-120823-022747, 2024, preprint: not peer 
reviewed.
Perin G, Bellan A, Bernardi A, Bezzo F, Morosinotto T. The potential of 
quantitative models to improve microalgae photosynthetic effi-
ciency. Physiol Plantarum. 2019:166(1):380–391. https://doi.org/10. 
1111/ppl.12915
Perin G, Bellan A, Michelberger T, Lyska D, Wakao S, Niyogi KK, 
Morosinotto T. Modulation of xanthophyll cycle impacts biomass 
productivity in the marine microalga nannochloropsis. Proc. Natl. 
3940 | The Plant Cell, 2024, Vol. 36, No. 10
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Acad. Sci. U S A. 2023:120(25):e2214119120. 10.1073/pnas. 
2214119120
Petrou K, Belgio E, Ruban AV. Ph sensitivity of chlorophyll fluores-
cence quenching is determined by the detergent/protein ratio 
and the state of LHCII aggregation. Biochim Biophys Acta. 2014: 
1837(9):1533–1539. https://doi.org/10.1016/j.bbabio.2013.11.018
Pi X, Zhao S, Wang W, Liu D, Xu C, Han G, Kuang T, Sui S-F, Shen J-R. 
The pigment-protein network of a diatom photosystem 
II-light-harvesting antenna supercomplex. Science. 2019:365-
(6452):eaax4406. https://doi.org/10.1126/science.aax4406
Prabhakar V, Lottgert T, Geimer S, Dormann P, Kruger S, 
Vijayakumar V, Schreiber L, Gobel C, Feussner K, Freussner I, 
et al. Phosphoenolpyruvate provision to plastids is essential for 
gametophyte and sporophyte development in Arabidopsis thali-
ana. Plant Cell. 2010:22(8):2594–2617. https://doi.org/10.1105/tpc. 
109.073171
Rast A, Schaffer M, Albert S, Wan W, Pfeffer S, Beck F, Plitzko JM, 
Nickelsen J, Engel BD. Biogenic regions of cyanobacterial thyla-
koids form contact sites with the plasma membrane. Nat Plants. 
2019:5(4):436–446. https://doi.org/10.1038/s41477-019-0399-7
Rea G, Lambreva M, Polticelli F, Bertalan I, Antonacci A, Pastorelli S, 
Damasso M, Johanningmeier U, Giardi MT. Directed evolution 
and in silico analysis of reaction centre proteins reveal molecular 
signatures of photosynthesis adaptation to radiation pressure. 
PLoS One. 2011:6(1):e16216. https://doi.org/10.1371/journal.pone. 
0016216
Roach T. LHCSR3-Type NPQ prevents photoinhibition and slowed 
growth under fluctuating light in Chlamydomonas reinhardtii. 
Plants. 2020:9(11):1604. https://doi.org/10.3390/plants9111604
Röding A, Boekema E, Büchel C. The structure of FCPb, a light- 
harvesting complex in the diatom Cyclotella meneghiniana. 
Photosynth Res. 2018:135(1–3):203–211. https://doi.org/10.1007/ 
s11120-016-0328-9
Rogov AG, Sukhanova EI, Uralskaya LA, Aliverdieva DA, Zvyagilskaya 
RA. Alternative oxidase: distribution, induction, properties, 
structure, regulation, and functions. Biochemistry (Mosc). 2014:79-
(13):1615–1634. https://doi.org/10.1134/S0006297914130112
Rosnow JJ, Edwards GE, Roalson EH. Positive selection of Kranz and 
non-Kranz C4 phosphoenolpyruvate carboxylase amino acids in 
Suaedoideae (Chenopodiaceae). J Exp Bot. 2014:65(13):3595–3607. 
https://doi.org/10.1093/jxb/eru053
Ruban AV, Johnson MP. Visualizing the dynamic structure of the 
plant photosynthetic membrane. Nat Plants. 2015:1(11):15161. 
https://doi.org/10.1038/nplants.2015.161
Ruban AV, Lavaud J, Rousseau B, Guglielmi G, Horton P, Etienne A-L. 
The super-excess energy dissipation in diatom algae: compara-
tive analysis with higher plants. Photosynth Res. 2004:82(2): 
165–175. https://doi.org/10.1007/s11120-004-1456-1
Sage RF, Zhu XG. Exploiting the engine of C(4) photosynthesis. J Exp 
Bot. 2011:62(9):2989–3000. https://doi.org/10.1093/jxb/err179
Saito S, Uozumi N. Guard cell membrane anion transport systems 
and their regulatory components: an elaborate mechanism con-
trolling stress-induced stomatal closure. Plants. 2019:8(1):9. 
https://doi.org/10.3390/plants8010009
Santana-Sánchez A, Nikkanen L, Werner E, Tóth G, Ermakova M, 
Kosourov S, Walter J, He M, Aro EM, Allahverdiyeva Y. Flv3A facil-
itates O2 photoreduction and affects H2 photoproduction inde-
pendently of Flv1A in diazotrophic Anabaena filaments. New 
Phytol. 2023:237(1):126–139. https://doi.org/10.1111/nph.18506
Santana-Sánchez A, Solymosi D, Mustila H, Bersanini L, Aro EM, 
Allahverdiyeva Y. Flavodiiron proteins 1-to-4 function in versa-
tile combinations in O2 photoreduction in cyanobacteria. Elife. 
2019:8:e45766. https://doi.org/10.7554/eLife.45766
Sarcina M, Bouzovitis N, Mullineaux CW. Mobilization of photosys-
tem II induced by intense red light in the cyanobacterium 
Synechococcus sp PCC7942. Plant Cell. 2006:18(2):457–464. https:// 
doi.org/10.1105/tpc.105.035808
Saroussi S, Sanz-Luque E, Kim RG, Grossman AR. Nutrient scaveng-
ing and energy management: acclimation responses in nitrogen 
and sulfur deprived Chlamydomonas. Curr Opin Plant Biol. 
2017:39:114–122. https://doi.org/10.1016/j.pbi.2017.06.002
Saroussi SI, Wittkopp TM, Grossman AR. The type II NADPH dehy-
drogenase facilitates cyclic electron flow, energy-dependent 
quenching, and chlororespiratory metabolism during acclima-
tion of Chlamydomonas reinhardtii to nitrogen deprivation. Plant 
Physiol. 2016:170(4):1975–1988. https://doi.org/10.1104/pp.15. 
02014
Schlau-Cohen GS, Yang H-Y, Krüger TPJ, Xu P, Gwizdala M, van 
Grondelle R, Croce R, Moerner WE. Single-molecule identification 
of quenched and unquenched states of LHCII. J Phys Chem Lett. 
2015:6(5):860–867. https://doi.org/10.1021/acs.jpclett.5b00034
Schneider AR, Geissler PR. Coexistence of fluid and crystalline 
phases of proteins in photosynthetic membranes. Biophys J. 
2013:105(5):1161–1170. https://doi.org/10.1016/j.bpj.2013.06.052
Schottkowski M, Peters M, Zhan Y, Rifai O, Zhang Y, Zerges W. 
Biogenic membranes of the chloroplast in Chlamydomonas rein-
hardtii. Proc. Natl. Acad. Sci. U S A. 2012:109(47):19286–19291. 
https://doi.org/10.1073/pnas.1209860109
Schubert H, Sagert S, Forster RM. Evaluation of the different levels of 
variability in the underwater light field of a shallow estuary. 
Helgoland Marine Research. 2001:55(1):12–22. https://doi.org/10. 
1007/s101520000064
Sejima T, Takagi D, Fukayama H, Makino A, Miyake C. Repetitive 
short-pulse light mainly inactivates photosystem I in sunflower 
leaves. Plant Cell Physiol. 2014:55(6):1184–1193. https://doi.org/10. 
1093/pcp/pcu061
Sello S, Meneghesso A, Alboresi A, Baldan B, Morosinotto T. Plant 
biodiversity and regulation of photosynthesis in the natural envi-
ronment. Planta. 2019:249(4):1217–1228. https://doi.org/10.1007/ 
s00425-018-03077-z
Sétif P, Shimakawa G, Krieger-Liszkay A, Miyake C. Identification 
of the electron donor to flavodiiron proteins in Synechocystis 
sp. PCC 6803 by in vivo spectroscopy. Biochim Biophys Acta 
Bioenerg. 2020:1861(10):148256. https://doi.org/10.1016/j.bbabio. 
2020.148256
Sharkey TD. The discovery of rubisco. J Exp Bot. 2022:74(2):510–519. 
https://doi.org/10.1093/jxb/erac254
Sharon I, Alperovitch A, Rohwer F, Haynes M, Glaser F, 
Atamna-Ismaeel N, Pinter RY, Partensky F, Koonin EV, Wolf YI, 
et al. Photosystem I gene cassettes are present in marine virus ge-
nomes. Nature. 2009:461(7261):258–262. https://doi.org/10.1038/ 
nature08284
Shi T, Bibby TS, Jiang L, Irwin AJ, Falkowski PG. Protein interactions 
limit the rate of evolution of photosynthetic genes in 
Cyanobacteria. Mol Biol Evol. 2005:22(11):2179–2189. https://doi. 
org/10.1093/molbev/msi216
Shikanai T, Endo T, Hashimoto T, Yamada Y, Asada K, Yokota A. 
Directed disruption of the tobacco ndhB gene impairs cyclic elec-
tron flow around photosystem I. Proc Natl Acad Sci U S A. 1998:95-
(16):9705–9709. https://doi.org/10.1073/pnas.95.16.9705
Shimakawa G, Miyake C. Changing frequency of fluctuating light re-
veals the molecular mechanism for P700 oxidation in plant 
leaves. Plant Direct. 2018:2(7):e00073. https://doi.org/10.1002/ 
pld3.73
Shimakawa G, Shaku K, Nishi A, Hayashi R, Yamamoto H, Sakamoto 
K, Makino A, Miyake C. FLAVODIIRON2 and FLAVODIIRON4 
Open questions in photosynthesis research | 3941
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
proteins mediate an oxygen-dependent alternative electron flow 
in Synechocystis sp. PCC 6803 under CO2-limited conditions. Plant 
Physiol. 2015:167(2):472–480. https://doi.org/10.1104/pp.114.249987
Shimoni E, Rav-Hon O, Ohad I, Brumfeld V, Reich Z. 
Three-dimensional organization of higher-plant chloroplast thy-
lakoid membranes revealed by electron tomography. Plant Cell. 
2005:17(9):2580–2586. https://doi.org/10.1105/tpc.105.035030
Smith WK, Berry ZC. Sunflecks? Tree Physiol. 2013:33(3):233–237. 
https://doi.org/10.1093/treephys/tpt005
Son M, Moya R, Pinnola A, Bassi R, Schlau-Cohen GS. Protein–protein 
interactions induce pH-dependent and zeaxanthin-independent 
photoprotection in the plant light-harvesting complex, LHCII. J 
Am Chem Soc. 2021:143(42):17577–17586. https://doi.org/10.1021/ 
jacs.1c07385
South PF, Cavanagh AP, Liu HW, Ort DR. Synthetic glycolate metab-
olism pathways stimulate crop growth and productivity in the 
field. Science. 2019:363(6422):eaat9077. https://doi.org/10.1126/ 
science.aat9077
Staehelin LA. Reversible particle movements associated with un-
stacking and restacking of chloroplast membranes in vitro. J 
Cell Biol. 1976:71(1):136–158. https://doi.org/10.1083/jcb.71.1.136
Standfuss J, van Scheltinga ACT, Lamborghini M, Kühlbrandt W. 
Mechanisms of photoprotection and nonphotochemical quench-
ing in pea light-harvesting complex at 2.5 Å resolution. EMBO J. 
2005:24(5):919–928. https://doi.org/10.1038/sj.emboj.7600585
Steen CJ, Burlacot A, Short AH, Niyogi KK, Fleming GR. Interplay be-
tween LHCSR proteins and state transitions governs the NPQ re-
sponse in intact cells of Chlamydomonas during light fluctuations. 
Plant Cell Environ. 2022:45(8):2428–2445. https://doi.org/10.1111/ 
pce.14372
Stirbet A, Lazár D, Guo Y, Govindjee G. Photosynthesis: basics, his-
tory and modelling. Ann Bot. 2020:126(4):511–537. https://doi. 
org/10.1093/aob/mcz171
Storti M, Alboresi A, Gerotto C, Aro E-M, Finazzi G, Morosinotto T. 
Role of cyclic and pseudo-cyclic electron transport in response 
to dynamic light changes in Physcomitrella patens. Plant Cell 
Environ. 2019:42(5):1590–1602. https://doi.org/10.1111/pce.13493
Streatfield SJ, Weber A, Kinsman EA, Hausler RE, Li J, 
Post-Beittenmiller D, Kaiser WM, Pyke KA, Flugge UI, Chory J. 
The phosphoenolpyruvate/phosphate translocator is required 
for phenolic metabolism, palisade cell development, and plastid- 
dependent nuclear gene expression. Plant Cell. 1999:11(9): 
1609–1622. https://doi.org/10.1105/tpc.11.9.1609
Suga M, Akita F, Hirata K, Ueno G, Murakami H, Nakajima Y, Shimizu 
T, Yamashita K, Yamamoto M, Ago H, et al. Native structure of 
photosystem II at 1.95 Å resolution viewed by femtosecond 
X-ray pulses. Nature. 2015:517(7532):99–103. https://doi.org/10. 
1038/nature13991
Sznee K, Dekker JP, Dame RT, van Roon H, Wuite GJL, Frese RN. 
Jumping mode atomic force microscopy on grana membranes 
from spinach. J Biol Chem. 2011:286(45):39164–39171. https://doi. 
org/10.1074/jbc.M111.284844
Taddei L, Stella GR, Rogato A, Bailleul B, Fortunato AE, Annunziata R, 
Sanges R, Thaler M, Lepetit B, Lavaud J, et al. Multisignal control 
of expression of the LHCX protein family in the marine diatom 
Phaeodactylum tricornutum. J Exp Bot. 2016:67(13):3939–3951. 
https://doi.org/10.1093/jxb/erw198
Tikkanen M, Grieco M, Kangasjärvi S, Aro E-M. Thylakoid protein 
phosphorylation in higher plant chloroplasts optimizes electron 
transfer under fluctuating light. Plant Physiol. 2010:152(2): 
723–735. https://doi.org/10.1104/pp.109.150250
Timm S, Mielewczik M, Florian A, Frankenbach S, Dreissen A, Hocken 
N, Fernie AR, Walter A, Bauwe H. High-to-low CO2 acclimation 
reveals plasticity of the photorespiratory pathway and indicates 
regulatory links to cellular metabolism of Arabidopsis. PLoS One. 
2012:7(8):e42809. https://doi.org/10.1371/journal.pone.0042809
Timm S, Woitschach F, Heise C, Hagemann M, Bauwe H. Faster re-
moval of 2-phosphoglycolate through photorespiration improves 
abiotic stress tolerance of Arabidopsis. Plants. 2019:8(12):563. 
https://doi.org/10.3390/plants8120563
Tiwari V, Matutes YA, Gardiner AT, Jansen TLC, Cogdell RJ, Ogilvie JP. 
Spatially-resolved fluorescence-detected two-dimensional elec-
tronic spectroscopy probes varying excitonic structure in photo-
synthetic bacteria. Nat Commun. 2018:9(1):4219. https://doi.org/ 
10.1038/s41467-018-06619-x
Tronconi MA, Hüdig M, Schranz ME, Maurino VG. Independent re-
cruitment of duplicated β-subunit-coding NAD-ME genes aided 
the evolution of C4 photosynthesis in Cleomaceae. Front Plant 
Sci. 2020:11:572080. https://doi.org/10.3389/fpls.2020.572080
Tsuchida Y, Furumoto T, Izumida A, Hata S, Izui K. 
Phosphoenolpyruvate carboxylase kinase involved in C(4) photo-
synthesis in Flaveria trinervia: cDNA cloning and characteriza-
tion. FEBS Lett. 2001:507(3):318–322. https://doi.org/10.1016/ 
S0014-5793(01)02994-5
Umena Y, Kawakami K, Shen J-R, Kamiya N. Crystal structure of 
oxygen-evolving photosystem II at a resolution of 1.9 Å. Nature. 
2011:473(7345):55–60. https://doi.org/10.1038/nature09913
van Grondelle R, Novoderezhkin VI. Energy transfer in photosynthe-
sis: experimental insights and quantitative models. Phys Chem 
Chem Phys. 2006:8(7):793–807. https://doi.org/10.1039/B514032C
Vicente JB, Carrondo MA, Teixeira M, Frazão C. Structural studies on 
flavodiiron proteins. Methods Enzymol. 2008:437:3–19. https://doi. 
org/10.1016/S0076-6879(07)37001-8
von Bismarck T, Korkmaz K, Ruß J, Skurk K, Kaiser E, Correa Galvis V, 
Cruz JA, Strand DD, Köhl K, Eirich J, et al. Light acclimation inter-
acts with thylakoid ion transport to govern the dynamics of pho-
tosynthesis in Arabidopsis. New Phytol. 2023:237(1):160–176. 
https://doi.org/10.1111/nph.18534
Walters RG, Ibrahim DG, Horton P, Kruger NJ. A mutant of 
Arabidopsis lacking the triose-phosphate/phosphate transloca-
tor reveals metabolic regulation of starch breakdown in the light. 
Plant Physiol. 2004:135(2):891–906. https://doi.org/10.1104/pp.104. 
040469
Wang D, Fiebig OC, Harris D, Toporik H, Ji Y, Chuang C, Nairat M, 
Tong AL, Ogren JI, Hart SM, et al. Elucidating interprotein energy 
transfer dynamics within the antenna network from purple bac-
teria. Proc Natl Acad Sci U S A. 2023a:120(28):e2220477120. https:// 
doi.org/10.1073/pnas.2220477120
Wang P, Frank A, Appel J, Boehm M, Strabel N, Nowaczyk MM, 
Schuhmann W, Conzuelo F, Gutekunst K. In vivo assembly of 
photosystem I-hydrogenase chimera for in vitro photoH2 produc-
tion. Adv Energy Mater. 2023b:13(14):2203232. https://doi.org/10. 
1002/aenm.202203232
Wang W, Yu L-J, Xu C, Tomizaki T, Zhao S, Umena Y, Chen X, Qin X, 
Xin Y, Suga M, et al. Structural basis for blue-green light harvest-
ing and energy dissipation in diatoms. Science. 2019:363(6427): 
eaav0365. https://doi.org/10.1126/science.aav0365
Wang W, Zhao S, Shen L, Li X, Tao Q, Xu C, Zhou C, Yang Y, Sang M, 
Han G, et al. Structural insights into photosystem II supercom-
plex and trimeric FCP antennae of a centric diatom Cyclotella me-
neghiniana. Nat Commun. 2023c:14(1):8164. https://doi.org/10. 
21203/rs.3.rs-3094926/v1
Wietrzynski W, Schaffer M, Tegunov D, Albert S, Kanazawa A, Plitzko 
JM, Baumeister W, Engel BD. Charting the native architecture of 
Chlamydomonas thylakoid membranes with single-molecule 
precision. Elife. 2020:9:e53740. https://doi.org/10.7554/eLife.53740
3942 | The Plant Cell, 2024, Vol. 36, No. 10
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024
Wood WHJ, MacGregor-Chatwin C, Barnett SFH, Mayneord GE, 
Huang X, Hobbs JK, Hunter CN, Johnson MP. Dynamic thylakoid 
stacking regulates the balance between linear and cyclic photo-
synthetic electron transfer. Nat Plants. 2018:4(2):116–127. 
https://doi.org/10.1038/s41477-017-0092-7
Wydrzynski T, Satoh S. Photosystem II: the light-driven water:plastoqui-
none oxidoreductase. Dordrecht (The Netherlands): Springer; 2005.
Xu C, Pi X, Huang Y, Han G, Chen X, Qin X, Huang G, Zhao S, Yang Y, 
Kuang T, et al. Structural basis for energy transfer in a huge dia-
tom PSI-FCPI supercomplex. Nat Commun. 2020:11(1):5081. 
https://doi.org/10.1038/s41467-020-18867-x
Yachandra VK, Sauer K, Klein MP. Manganese cluster in photosyn-
thesis: where plants oxidize water to dioxygen. Chem Rev. 
1996:96(7):2927–2950. https://doi.org/10.1021/cr950052k
Yamamoto H, Peng L, Fukao Y, Shikanai T. An Src homology 3 
domain-like fold protein forms a ferredoxin binding site for the 
chloroplast NADH dehydrogenase-like complex in Arabidopsis. 
Plant Cell. 2011:23(4):1480–1493. https://doi.org/10.1105/tpc.110. 
080291
Yamamoto H, Shikanai T. PGR5-Dependent cyclic electron flow pro-
tects photosystem I under fluctuating light at donor and acceptor 
sides. Plant Physiol. 2018:179(2):588–600. https://doi.org/10.1104/ 
pp.18.01343
Yamamoto H, Takahashi S, Badger MR, Shikanai T. Artificial remod-
elling of alternative electron flow by flavodiiron proteins in 
Arabidopsis. Nat Plants. 2016:2(3):16012. https://doi.org/10.1038/ 
nplants.2016.12
Yano J, Yachandra VK. Mn4Ca cluster in photosynthesis: where and 
how water is oxidized to dioxygen. Chem Rev. 2014:114(8): 
4175–4205. https://doi.org/10.1021/cr4004874
Yokochi Y, Yoshida K, Hahn F, Miyagi A, Wakabayashi K-i, 
Kawai-Yamada M, Weber APM, Hisabori T. Redox regulation of 
NADP-malate dehydrogenase is vital for land plants under fluc-
tuating light environment. Proc Natl Acad Sci U S A. 2021:118(6): 
e2016903118. https://doi.org/10.1073/pnas.2016903118
You X, Zhang X, Cheng J, Xiao Y, Ma J, Sun S, Zhang X, Wang HW, Sui 
SF. In situ structure of the red algal phycobilisome–PSII–PSI–LHC 
megacomplex. Nature. 2023:616(7955):199–206. https://doi.org/ 
10.1038/s41586-023-05831-0
Zabret J, Bohn S, Schuller SK, Arnolds O, Möller M, Meier-Credo J, 
Liauw P, Chan A, Tajkhorshid E, Langer JD, et al. Structural in-
sights into photosystem II assembly. Nat Plants. 2021:7(4): 
524–538. https://doi.org/10.1038/s41477-021-00895-0
Zelitch I, Walker DA. The role of glycolic acid metabolism in opening 
of leaf stomata. Plant Physiol. 1964:39(5):856–862. https://doi.org/ 
10.1104/pp.39.5.856
Zeno G, Gomez RL, Caferri R, Stuttmann J, Dall’Osto, L, Bassi R. 
Thylakoid grana stacking revealed by multiplex genome editing 
of LHCII encoding genes. bioRxiv 474624, https://doi.org/10. 
1101/2021.12.31.474624, 1 January 2022, preprint: not peer 
reviewed.
Open questions in photosynthesis research | 3943
D
ow
nloaded from
 https://academ
ic.oup.com
/plcell/article/36/10/3914/7718004 by Library of M
edical Faculty user on 10 O
ctober 2024