REDOX REGULATION OF CYCLIC ELECTRON FLOW AROUND
PHOTOSYSTEM I
By
Deserah Dawn Strand

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Plant Biology—Doctor of Philosophy
2014
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ABSTRACT
REDOX REGULATION OF CYCLIC ELECTRON FLOW AROUND
PHOTOSYSTEM I
By
Deserah Dawn Strand
The proton coupled electron transfer of the light reactions of photosynthesis
supply ATP and reducing power to metabolic processes within the chloroplast and,
ultimately, the organism. The textbook pathway of electron transfer or ‘Z-scheme’ of
photosynthesis supplies a fixed ratio of ATP and reducing power (i.e. reduced ferredoxin
and NAD(P)H), however, the downstream metabolic demands require a highly dynamic
ratio. This poses a problem when downstream metabolic demands lead to a deficit of
ATP, a buildup of reducing equivalents may occur in the stroma, which may then lead to
the buildup of reactive oxygen species (ROS). To balance the output of the light
reactions, the chloroplast employs several mechanisms to either shunt electrons out of the
system, or by redirecting electrons away from downstream metabolism and back into the
plastoquinone pool to generate additional ATP in a process termed cyclic electron flow
around photosystem I (CEF).
CEF has been proposed to be catalyzed by multiple pathways. In plants, this
includes the antimycin A sensitive ferredoxin quinone reductase (FQR) and the
respiratory Complex I analog, the NADPH dehydrogenase complex (NDH). We found
these pathways are differentially regulated by redox status, with the FQR more active in a
reducing environment (Em = -306 mV), while the NDH is activated in response to ROS.
We further found that the kinetics of activation for the two pathways were different, with
the FQR rapidly activated (seconds to minutes) and the NDH activated more slowly

(minutes). Additionally, we have evidence that the NDH is a proton pumping
plastoquinone reductase. This observation indicates the two pathways contribute different
levels of pmf to offset an ATP deficit. We propose CEF pathways are differentially
regulated in response to chloroplast redox state. The FQR is rapidly activated when
reducing equivalents in the stroma accumulate in response to a short-term or rapidly
fluctuating ATP deficit. When the FQR fails to restore homeostasis and the ATP deficit
leads to ROS formation, the NDH complex is activated as a more robust mechanism of
pmf generation via CEF.

Copyright by
DESERAH DAWN STRAND
2014

This dissertation is dedicated to Dr. Virginia Lee, a truly inspiring person.

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ACKNOWLEDGEMENTS

I’ve received much academic and personal support over the course of my graduate career.
I would like to thank the entirety of the Kramer lab, both at Michigan State University and
Washington State University, for their unending willingness to help me through whatever I
needed from them. My advisor, Dr. David Kramer, has been an amazing source of knowledge
and inspiration. I have been challenged almost everyday working with the Kramer lab. I’d also
like to specifically thank Dr. Jeffrey Cruz for the hours of technical (and oftentimes emotional)
support he has given me. Robert Zegarac has been an enormous help by providing his technical
expertise. Dr. Kaori Kohzuma was an excellent resource and the best office mate I could have
asked for. Dr. Nicholas Fisher has been both inspiring and insightful throughout his time at
MSU. Dr. Sean Weiss has also been very helpful and encouraging in many aspects of my more
difficult projects. Dr. Ben Lucker and Chris Hall were more than willing to help me figure out
how to grow and experiment on Chlamydomonas, and while none of this work made it into this
dissertation, they have been of valuable assistance along the way.
In addition to Chris Hall, I’ve had the great opportunity to work with an excellent group
of students at both MSU and WSU. At MSU, L. Ruby Carrillo has been a great sounding board
for ideas and an overall joy to work with. I hope we have the opportunity to continue to work
together in the future. A recent addition to the Kramer lab, Geoff Davis, has also been wonderful
to work with. At WSU, Dr. Stefanie Tietz and Amelia Barhanovich helped me get through the
transition to, not only graduate school, but also to a new state, and Dr. Aaron Livingston helped
me (smoothly) transition into the chaos of the Kramer lab. Dr. Elisabeth Ostendorf was an

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excellent roommate and friend, first as a visiting student at WSU, and currently as a post-doc at
MSU.
I would also like to thank all the undergraduate research assistants I had the chance to
work with throughout the years. Ben Wolf, Brendan Johnson, and Heather Enlow were
particularly amazing to work with. I couldn’t have done so much without them.
I would also like to thank Dr. Dale Vogelien, at Kennesaw State University, for the
excellent experiences as an undergraduate that led me to trying for a career in plant research. I
learned so much from, and was inspired everyday, working with her. Also at KSU, I am very
grateful for the opportunities to work with Dale Zaborowski, Dr. Marina Koether and Dr.
Thomas McElroy. I believe that these early experiences helped shape me as a scientist.
Finally, I would like to thank my friends and family for their ongoing support in everything I do.
Ariel Smith (now Rosshirt), has always been there to talk to, whether I was frustrated or
homesick. My parents have been of constant support throughout my years at MSU and I am very
grateful for this. I would also like to thank my brother, Jason, for being entirely positive and
encouraging.

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TABLE OF CONTENTS

LIST OF FIGURES

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Chapter 1 Control of non-photochemical exciton quenching by the proton circuit
of photosynthesis
1.1 Abstract
1.2 Introduction
1.2.A The electron and proton circuits of photosynthesis
1.2.B The proton circuit is a central regulation of photosynthesis
1.3 Type I flexibility mechanisms: NPQ and balancing of the chloroplast
energy budget
1.3.A Malate valve
1.3.B Mehler peroxidase reaction (MPR)
1.3.C Plastid terminal oxidase (PTOX)
1.3.D Exchange of ATP/ADP+Pi between the stroma and cytosol
1.3.E Cyclic electron flow (CEF) around photosystem I
1.3.E.1 CEF pathways
1.3.E.1.A Chloroplast NDH pathway
1.3.E.1.B Supercomplex pathways
1.3.E.1.C The ferredoxin-plastoquinone reductase (FQR)
pathways
1.3.E.1.D NDA2 pathway
1.3.E.2 Regulation of CEF
1.4 Type II flexibility mechanisms: regulation of pmf partitioning and
ATP synthase activity and the consequences for NPQ
1.4.A The chloroplast ATP synthase is a central regulator of
photosynthesis
1.4.B Regulation of pmf partitioning
1.5 Concluding remarks
1.6 Acknowledgements
Chapter 2 Redox regulation of the antimycin A sensitive pathway of cyclic electron
flow around photosystem I in higher plant thylakoids
2.1 Abstract
2.2 Introduction
2.3 Results
2.3.A Effect of reductant on CEF activity in vitro
2.3.B Dependency of Fd on CEF activity
2.3.C The redox dependence of CEF activation by DTT
2.4 Discussion
2.4.A FQR-mediated CEF is active under reducing conditions in vitro
2.4.B FQR-mediated CEF may involve a regulatory thiol

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2.4.C FQR-mediated CEF is sensitive to thylakoid isolation
conditions
2.4.D Implication for regulation in vivo, two pathways of CEF
2.5 Methods
2.5.A Chloroplast isolation
2.5.B Oxidation and reduction of dithiothreitol (DTT)
2.5.C Spectroscopic measurements
2.5.C.1 Acridine fluorescence quenching
2.5.C.2 Absorbance changes at 703 nm
2.6 Author contributions
2.7 Acknowledgements
Chapter 3 Activation of cyclic electron flow around photosystem I by hydrogen
peroxide in vivo
3.1 Abstract
3.2 Introduction
3.3 Results
3.3.A H2O2 accumulation in the high cyclic mutant hcef1

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3.3.B Effects of H2O2 production by plants expressing glycolate
oxidase in the chloroplast
3.3.C Kinetics of CEF induction upon activation of H2O2
production in GO5
3.3.D Induction of CEF by infiltration of leaves with H2O2
3.4 Discussion
3.4.A CEF correlates with, and is induced by, H2O2 production
in vivo
3.4.B H2O2 activates the antimycin A insensitive pathway

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3.4.C A regulatory role for H2O2 in activation of CEF?
3.5 Methods
3.5.A Plant material and growth conditions
3.5.B In vivo spectroscopic assays
3.5.C Infiltration of leaves with antimycin A and H2O2
3.5.D Manipulation of gas concentrations
3.5.E Measurement of H2O2 production in leaves
3.5.F Protein extraction and western blot
3.5.G Quantifications of CEF
3.5.H Statistical analysis
3.6 Author contributions
3.7 Acknowledgements

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Chapter 4 Uncoordinated expression of chloroplast proteins lead to H2O2
accumulation and activation of cyclic electron flow around photosystem I
4.1 Abstract
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4.2 Introduction
4.3 Results
4.3.A Genetic selection of hcef mutants
4.3.B Growth of hcef2
4.3.C Photosynthetic electron transfer properties of hcef2 compared
to Col-0: responses of the photosynthetic electron transport to
the hcef2 mutation
4.3.D Responses of the photosynthetic proton circuit of hcef2
4.3.E Estimates of CEF in hcef2
4.3.F Antimycin A infiltration of hcef2
4.3.G Identification of the genetic locus of hcef2 as TADA1
4.3.H Translational defects in the chloroplast lead to increases in CEF
4.3.I Mass spectrometry proteomics
4.3.J Flash induced relaxation kinetics in dark-adapted leaves of hcef2
4.3.K hcef2 and related mutants show elevated H2O2 production
4.4 Discussion
4.4.A Disruption of protein translation in hcef2 leads to accumulation
of H2O2 and activation of CEF involving the chloroplast NDH
complex
4.4.B The hcef2 mutantion disrupts chloroplast translation and
leads to mis-assembly of chloroplast protein complexes
4.4.C Modifying chloroplast translation leads to accumulation of
H2O2, a likely activator of CEF
4.5 Methods
4.5.A Plant materials and growth
4.5.B Isolation of hcef mutants
4.5.C 77K fluorescence spectroscopy
4.5.D In vivo spectroscopy
4.5.E Map-based cloning of hcef2
4.5.F CAPS marker for hcef2 genotype
4.5.G Infiltrations
4.5.H H2O2 quantification
4.5.I Determination of protein changes in hcef2
4.6 Author contributions
4.7 Acknowledgements
Chapter 5 Evidence for a proton pumping plastidic NAD(P)H dehydrogenase
complex involved in cyclic electron flow around photosystem I
5.1 Abstract
5.2 Introduction
5.3 Results
5.4 Discussion
5.4.A CEF involves a proton pump
5.4.B A proton pumping NDH would be reversible
5.5 Methods

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5.5.A Plant materials and growth conditions
5.5.B In vivo spectroscopy
5.5.C Quantification of in vitro ATP production
5.6 Author contributions
5.7 Acknowledgements

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Chapter 6 Conclusions and future directions
6.1 Abstract
6.2 Conclusions
6.3 Future directions

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APPENDICES
Appendix 1 Supplemental methods for chapter 4
Appendix 2 Towards a more precise measure of electron transfer rates
through photosystem II

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REFERENCES

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LIST OF FIGURES

Figure 1.1 The light-dependant electron and proton transfer reactions of
photosynthesis .

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Figure 1.2 The malate valve.

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Figure 1.3 The Mehler peroxidase reaction.

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Figure 1.4 The plastid terminal oxidase (PTOX).

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Figure 1.5 Proposed routes of cyclic electron flow around photosystem I.

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Figure 1.6 Proposed mechanisms of changes in pmf partitioning.

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Figure 2.1 Light induced formation of pmf monitored by acridine fluorescence
-2 -1
quenching upon illumination with 200 µmol photons m s photosynthetically
active radiation.

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Figure 2.2 Light induced P700 redox changes.

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Figure 2.3 Irreversible loss of CEF in thylakoids isolated in the absence of DTT.

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Figure 2.4 Dependence of CEF on Fd in vitro.

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Figure 2.5 Redox titration of CEF activity under varying concentrations of
DTTOX/2DTTRED.

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Figure 2.6 Rapid reversibility of CEF.

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Figure 2.7. A comprehensive model of CEF activation in vivo.

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Figure 3.1 Relative H2O2 accumulation in hcef1.

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Figure 3.2 Relative H2O2 accumulation in GO5.

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Figure 3.3 Effects of chloroplast targeted glycolate oxidase expression on
photosynthesis .

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Figure 3.4 Increased CEF in GO5.

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Figure 3.5 Increased dark reduction of the PQ/PQH2 pool in GO5.

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Figure 3.6 Increased CEF in mutants co-transformed with glycolate oxidase (GO)
and malate synthase (MS) targeted to the chloroplast.

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Figure 3.7 Induction kinetics of H2O2 activated CEF.

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Figure 3.8 Increased dark reduction of the PQ/PQH2 pool in leaves infiltrated with
0.1% H2O2 for 3 hours.

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Figure 3.9 Model of CEF activation in response to H2O2.

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Figure 4.1 Growth phenotype of hcef2 at 24 days.

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Figure 4.2 Effects of hcef2 on photosynthetic properties.

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Figure 4.3 Effects of hcef2 on the photosynthetic proton circuit.

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Figure 4.4 Activation state of CEF in Col-0 and hcef2.

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Figure 4.5 Kinetics of postillumination fluorescence rises in Col-0 and hcef2.

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Figure 4.6 Antimycin A insensitivity of CEF in hcef2.

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Figure 4.7 Introduction of a stop codon into the TADA1 locus in hcef2.

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Figure 4.8 Elevated CEF in tada1.

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Figure 4.9 TADA1 complementation of hcef2.

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Figure 4.10 Elevated CEF in the ribosomal mutants prsp3 and rps17.

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Figure 4.11 Changes in the thylakoid proteome of hcef2.

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Figure 4.12 Flash induced relaxation kinetics of the electrochromic shift in Col-0
(black) and hcef2 (gray) leaves infiltrated with 100 µM methyl viologen.

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Figure 4.13 Relative accumulation of H2O2 in Col-0 and mutant lines.

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5.1 Residues essential for proton pumping are conserved in the NDH
membrane fraction.

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Figure 5.2 Evidence for involvement of a proton pump in CEF.

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Figure 5.3 Directionality of NDH.

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Figure 5.4 Hypothetical model of the reversible reactions of NDH.

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Figure A2.1 Representative experimental data for the calculation of [PSI].

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Figure A2.2 Representative experimental data for the calculation of E.

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Figure A2.3 Decreased [PSI] in hcef1.

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Figure A2.4 Altered [PSII]/[PSI] in hcef1.

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Chapter 1
Control of non-photochemical exciton quenching by the proton circuit of photosynthesis1
Deserah D. Strand and David M. Kramer

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1
This chapter is modified from Strand, D.D., and Kramer, D.M. (2014) Control of NonPhotochemical Quenching by the Proton Circuit of Photosynthesis. In Advances in
Photosynthesis and Respiration, Vol. 40, Non-Photochemical Quenching and Energy Dissipation
in Plants, Algae and Cyanobacteria Demmig-Adams, B., Garab, G., Adams III, W., Govindjee
(Eds) Springer Netherlands DOI:10.1007/978-94-017-9032-1 ISBN 978-94-017-9031-4
1

1.1 Abstract
This review discusses our current understanding of the chloroplast proton circuit, i.e.
those reactions that involve the storage and utilization of light energy in the transfer of protons,
and its importance for regulating photosynthetic light-capture/electron-transfer reactions. The
photosynthetic machinery of plants is finely tuned to balance the needs for efficient light capture
with an avoidance of photodamage by regulating the capture of light energy, via thermal
dissipation of excess excitation energy (assessed from non-photochemical quenching, NPQ, of
chlorophyll a fluorescence) by regulating light-driven electron transfer processes. In addition to
driving ATP synthesis at the chloroplast ATP synthase, the thylakoid electrochemical gradient of
protons or proton-motive force (pmf) plays a central role in regulating NPQ. The transthylakoid
proton concentration gradient (!pH) component of pmf triggers the “energy-dependent”, or qE
component of NPQ, which protects photosystem II from photodamage and regulates electron
transfer through the cytochrome b6f complex, thereby preventing damage to photosystem I. The
extent and mode of storage in !pH and !" of pmf are regulated by several processes that
respond to the metabolic, or physiological, state of the organism. The extent of pmf is determined
by proton influx (via linear and alternative electron flows) into the thylakoid lumen, and proton
efflux through the chloroplast ATP synthase. Both processes are modulated by, or responsive to,
environmental conditions and resulting metabolic fluctuations. Proton influx is controlled by
linear electron flow and a series of alternative electron flow pathways, possibly including cyclic
electron flow around photosystem I, the Mehler peroxidase reaction (or water-water cycle), and
oxidation of plastoquinol by the plastid terminal oxidase. The fraction of pmf stored as !pH is
also regulated by plastidic ionic strength or luminal buffering capacity, altering the sensitivity of

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pH-dependent processes to pmf. The integrated regulation of these processes is an open, active
area of research.
1.2 Introduction
Sunlight, harnessed in photosynthesis, is the energy source for most life on our planet.
Energy from absorbed sunlight presents a challenge for photosynthetic organisms because within
the light reactions of photosynthesis, highly reactive redox intermediates are formed that can
generate harmful reactive oxygen species (ROS). Thus, plants must regulate photosynthetic light
harvesting and electron transfer to avoid photodamage, especially when light input exceeds the
capacity of photosynthesis to safely process it (1). To meet this need, essentially every process in
photosynthesis is regulated to balance minimization of deleterious side reactions with the need
for efficient capture of solar energy (2, 3). This regulation is critical for efficient energy
conversion, since most of the energy captured by an ecosystem is lost within the light reactions
of photosynthesis, much of it dissipated in photoprotective processes, the level of which can be
estimated from the non-photochemical quenching (NPQ) of chlorophyll a fluorescence (4–7).
This is a critical point: even though the rate-limiting steps in photosynthesis occur in biochemical
processes that follow the initial storage of energy by the light reactions, these limitations
inevitably result in the loss of energy by one of two mechanisms: backup of electrons and
protons resulting in long-lived excitation states (and potential ROS formation), or activation of
processes that “dump” or dissipate energy in excess of that needed to sustain the downstream
processes. As discussed below, the former mechanism can result in the buildup of deleterious
intermediates while the second prevents their generation, but, if not tightly regulated, could lead
to excessive energy loss.

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Over developmental time scales, this regulation can occur by altering such processes as
gene expression, protein accumulation, and the synthesis of pigments and antioxidants (3, 8).
However, plants must also respond to rapid fluctuations in environmental conditions such as
light and temperature, requiring equally rapid regulation at the level of the photosynthetic
machinery itself (1, 9).
One of the major mechanisms for this regulation is NPQ of antenna excitation energy, a
key component of which, qE (“energy dependent” quenching), is triggered by acidification of the
thylakoid lumen caused by the transthylakoid proton concentration gradient, or !pH component
of thylakoid proton-motive force (pmf) (9, 10). The pmf is generated by the light-induced
electron transfer reactions and, in turn, drives photophosphorylation of ADP to ATP. In this
paper, we review our understanding of how the process of activating qE is intimately integrated
into the entire energy storage system to balance energy capture and storage.
1.2.A The electron and proton circuits of photosynthesis
The light reactions of oxygenic photosynthesis consist of two highly integrated energystoring circuits. The “electron circuit” stores energy through a series of light-driven electron
transfer reactions, via the linear electron flow (LEF) pathway (Figure 1.1) (3). Light energy is
captured by pigments in light-harvesting complexes and shunted to a special subset of
chlorophylls in photosystem I (PS I) and photosystem II (PS II) (11–13). The photochemically
excited chlorophylls in PS II extract electrons from water and transfer them through an electron
transfer chain, reducing plastoquinone (PQ) to plastoquinol (PQH2), then to the cytochrome b6f
complex (b6f), plastocyanin, and finally to PS I. An additional photon of light energy is

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Figure 1.1 The light-dependent electron and proton transfer reactions of photosynthesis.
Electrons are extracted from water and passed through an electron transport chain. Protons are
translocated into the lumen generating a pmf to drive ATP synthesis. Stroma (S), Lumen (L),
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Photosystem I (PS I), Photosystem II (PS II), Plastoquinone (PQ), Ferredoxin-NADP reductase
(FNR), Ferredoxin (Fd), Plastocyanin (PC), the cytochrome b6f complex (b6f).

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introduced at PS I, driving electron transfer to ferredoxin (Fd) and NADP , which are then used
to drive reductive processes in the cell.
The “proton circuit” is energetically and mechanistically coupled to the electron circuit to
store energy in an electrochemical gradient of protons between the stroma and thylakoid lumen
across the thylakoid membrane, or pmf, which drives the synthesis of ATP through the ATP
synthase that spans the thylakoid membrane. Protons are released into the thylakoid lumen
during water oxidation at PS II. Additional protons are translocated via the Q-cycle, an
enzymatic redox loop catalyzed by the b6f complex and related complexes involving the
oxidation and reduction of plastoquinones and subsequent electron and proton transfer reactions
that results in proton translocation across the thylakoid membrane (14, 15).
1.2.B The proton circuit is a central regulator of photosynthesis
When light absorption exceeds the capacity of the downstream reactions of
photosynthesis, reactive intermediates may accumulate potentially leading to photoinhibition and
photodamage, although the relationship between reactive intermediates and photoinhibition is
still controversial [reviewed in (16)]. Plants have multiple mechanisms of avoidance and
dissipation of excess light, including activation of processes underlying NPQ, changes in leaf
orientation, chloroplast movement, rearrangement of the photosynthetic machinery, and changes
in photosynthetic gene expression (8, 17–19).
This chapter focuses on how the activation of a process reflected in NPQ and other
processes that regulate light capture and electron flow interact with the proton circuit of
photosynthesis. We concentrate mainly on these processes in higher plants and, where
appropriate, in algae. The case of cyanobacteria is not covered here because, in these organisms,
NPQ is controlled by factors not directly related to pmf (20, 21).
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Much of the dynamic regulation of photosynthesis directly involves the thylakoid proton
circuit, which modulates light capture, photoprotection, and electron transfer allowing the light
reactions to respond to changes in the physiology of the organism. Chloroplasts are thought to
protect themselves from photodamage by dissipating excess light energy, as reflected in NPQ (1,
22). The major, rapidly reversible component of NPQ in plants is termed qE (for energydependent quenching), and is triggered by the !pH component of pmf, which acidifies the
lumen, thereby activating violaxanthin deepoxidase, catalyzing the conversion of violaxanthin to
antheraxanthin and zeaxanthin (23), and protonating the antenna-associated protein PsbS (10, 22,
24).
Photosynthetic electron flow is regulated at the b6f complex by acidification of the lumen.
As a result of the buildup of the !pH component of pmf, plastoquinol oxidation is slowed at the
Qo site (quinol oxidation) (25, 26). This prevents accumulation of electrons on highly reducing
components of the acceptor side of PS I, which can lead to generation of reactive oxygen species
(ROS) and/or affect redox regulation of metabolism.
In extreme cases, lumen acidification can sensitize PS II to light-induced destabilization
(27, 28). There is evidence that this can occur in vivo under low CO2 or environmental stresses,
such as drought (26). This is often called photoinhibition. Because it produces inactive PS II,
which quenches excitation energy and decreases electron transfer, photoinhibition may represent
a regulatory “tactical retreat" to prevent overexcitation and damage to DNA or other cellular
components (29–31).
The proton circuit is, in turn, modulated by secondary regulatory mechanisms,
particularly at the ATP synthase, allowing the light reactions to respond to changes in the
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physiology of the organism. The activity of the ATP synthase is downregulated to retard proton
efflux from the thylakoid lumen, increasing pmf and leading to down-regulation of light capture
by qE and regulation of electron transfer reactions at the b6f complex (9, 25, 32–35). The fraction
of pmf stored as !pH and electric field (!") affects the relationship between pmf and lumen
acidification, further adjusting the pH-dependent responses underlying NPQ (33, 36). In
addition, the responses of qE to lumen pH can be modulated by altering the expression of qErelated proteins (10, 37, 38).
These “flexibility mechanisms”, which may adjust pmf-dependent NPQ responses to
physiological and metabolic changes, fall into two categories (2, 25). “Type I” mechanisms
adjust the “qE response” (defined as the activation state of qE as a function of LEF) by altering
light-driven proton-transfer reactions via activation of alternative cyclic electron transfer
processes and, because proton efflux from the lumen is generally coupled to ATP synthesis,
affect the ratio of ATP/NADPH arising from photosynthetic electron transport. “Type II”
flexibility mechanisms, on the other hand, adjust the qE response without altering the
ATP/NADPH balance. The distinction between Type I and Type II mechanisms is critical
because, despite both having an effect on qE, they have very different consequences for
downstream metabolism, i.e., the requirement for the ATP/NADPH ratio to precisely match that
required by downstream reactions may pose substantial constraints on the activation of Type I
mechanisms.
1.3 Type I flexibility mechanisms: NPQ and balancing of the chloroplast energy budget
Proton and electron transfer reactions in photosynthetic complexes are tightly coupled
and consumption of photosynthetic products must thus match production to prevent buildup of
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reactive intermediates (2, 3, 39). The ratio of ATP/NADPH from LEF is thought to be fixed at
1.3, approximately 13% less than the 1.5 required for the Calvin-Benson-Bassham (CBB) cycle
and other metabolic processes in the chloroplast [reviewed in (2)]. The ATP and NADPH
consumption within the chloroplast is also dynamic and dependent on the activation of multiple
processes and thus requires rapidly adjustable plasticity in the relative output of ATP and
NADPH. At least five processes have been proposed to remedy a deficit of ATP in the
chloroplast (2, 3, 40, 41), as discussed in the following sections.
1.3.A Malate valve
The “malate valve” (Figure 1.2) (41) involves shuttling of reducing equivalents from the
chloroplast to other cellular compartments. NADP-malic enzyme (MDH) uses NADPH to reduce
oxaloacetate to malate. The malate can then be exported from the chloroplast to the mitochondria
to generate ATP from respiration. In this process, ATP production is maintained while NADPH
is only transiently reduced, allowing it to help balance the chloroplast ATP/NADPH ratio. MDH
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is inhibited by NADP , suggesting a control mechanism involving NADPH levels, which would
be consistent with an involvement in balancing production of ATP and NADPH in the
chloroplast. On the other hand, it has been argued that the activity of the malate valve is too low
to account for substantial ATP/NADPH rebalancing under permissive conditions, i.e., conditions
like those in the laboratory, where loss of productivity to environmental stress is minimal (42,
43), although there is also evidence that malate valve activity is increased under prolonged stress
where the ratio of ATP/NADPH consumption may be higher (41). Supporting this evidence,
Hebbelman et al. (44) found upregulation of antioxidant pathways in mutants lacking MDH
when exposed to high light, suggesting that the malate valve may play expanded roles under nonlaboratory conditions.
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Figure 1.2 The malate valve. NADPH is consumed in the production of malate from
oxaloacetate (OAA) via NADP-malate dehydrogenase (MDH). The malate is then exported from
the chloroplast.

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1.3.B Mehler peroxidase reaction (MPR)
In the MPR, also referred to as the water-water cycle (Figure 1.3), electrons from LEF are
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shunted away from NADP reduction and instead reduce O2 to superoxide at PS I (45–47).
Superoxide is then dismutated to H2O2 and O2 via superoxide dismutase and then H2O2 is
reduced to H2O via ascorbate peroxidase. This leads to production of ATP via LEF-coupled
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accumulation of protons in the thylakoid lumen, without net reduction of NADP (48). The MPR
appears to contribute significantly to ATP synthesis and photoprotection in cyanobacteria and
bryophytes, but appears to have limited in vivo capacity in vascular plants and algae under
permissive steady state conditions (42, 49–54). It has been reported that MPR may have an
important role during activation of photosynthesis to prevent over-reduction of the electron
transport chain under excess light (46, 55, 56).
1.3.C Plastid terminal oxidase (PTOX)
Chloroplasts also contain an alternate oxidase (PTOX) that oxidizes PQH2 and reduces
O2 to H2O (Figure 1.4) (57). Electron flow from PS II through PTOX could generate ATP from
proton accumulation at PS II; however, its capacity would be limited by the fact that only one
proton is expected to be deposited into the lumen per electron transferred from PS II through
PTOX. In vascular plants, PTOX is important in developing plastids (58) and some alpine plants
(59, 60), although its contribution to ATP/NADPH balance in steady-state photosynthesis in
most species is probably small (2, 3). It has recently been proposed that PTOX works in
conjunction with the chloroplast NADPH:plastoquinone oxidoreductase (NDH, see below) to

11

Figure 1.3 The Mehler peroxidase reaction. Electrons from PS I reduce O2 to superoxide (O2
), which is dismutated to H2O2 via superoxide dismutase (SOD), and then reduced to H2O and
O2 via ascorbate peroxidase (APX), oxidizing ascorbate, or ascorbic acid (AsA), to
monodehydroascorbate (MDA).

Figure 1.4 The plastid terminal oxidase (PTOX). Electrons from PS II reduce plastoquinone
(PQ), which is then oxidized via PTOX, bypassing the Q-cycle. Proton translocation into the
lumen is only possible via the plastoquinone reductases (PS II in this figure).
tune the redox state of the photosynthetic electron transport chain, and possibly acts as an
electron sink under high-light conditions in tomato (60, 61).

12

-

1.3.D Exchange of ATP and ADP+Pi between the stroma and cytosol
The ATP budget could also be balanced by direct exchange of ATP and ADP + inorganic
phosphate (Pi) among chloroplast, cytosol, and mitochondria. Early suggestions that chloroplasts
directly exchange ATP and ADP+Pi with the cytosol were questioned by observations of slow
ATP and ADP exchange in isolated chloroplasts (62, 63), leading to the generally accepted view
that the chloroplast ATP budget is “self-contained”. However, this has not been rigorously tested
under physiologically relevant conditions, opening up the possibility that ATP and ADP+Pi
exchange may, in fact, be sufficiently rapid to allow highly flexible energy balancing.
1.3.E Cyclic electron flow (CEF) around PS I
CEF is strongly implicated in the ATP/NADPH balance (2, 3, 64–66). CEF involves
photochemistry in PS I, but not PS II, resulting in oxidation of plastocyanin and reduction of Fd.
Electrons from PS I are transferred back into the PQ pool in a ferredoxin-dependent process
involving a plastoquinone reductase (PQR). Following its formation by PQR, PQH2 is oxidized
by the b6f complex via the Q-cycle (14, 15), resulting in proton translocation into the thylakoid
lumen that generates pmf. This pmf is then used to drive ATP synthesis. The electrons are passed
back to PS I via plastocyanin. This process results in ATP formation, with no net NADPH
production, altering the ATP/NADPH stoichiometry formed by the light reactions.
CEF appears to be minimally activated under non-stressed conditions in C3 plants, when
LEF nearly meets the ATP required for chloroplast metabolism (9, 35, 67–70). However, CEF is
upregulated when high ATP demand is greater, e.g., under environmental stress (52, 59, 71, 72),
when carbon-concentrating mechanisms or C4 photosynthesis are engaged (73, 74), or during
13

induction of photosynthesis when dark-adapted plants are suddenly exposed to light (75, 76).
CEF may also play a regulatory role via acidification of the thylakoid lumen (25, 66, 77). These
ATP/NADPH balancing and regulatory roles for CEF are not independent, since ATP/NADPH
balance is critical for maintaining pmf, and thereby regulating qE or electron flux through the b6f
complex. However, CEF is clearly not essential for qE since qE can be observed in the absence of
CEF (35), when other processes, including the regulation of the ATP synthase and partitioning of
pmf into !pH and !" components, can account for regulation of qE responses (25, 32, 33, 78).
Several labs have isolated mutants, in Arabidopsis and tobacco, exhibiting high rates of
CEF (hcef). Characterization of these mutants has established that CEF can run at high rates even
in C3 plants, but must also be highly regulated in the wild type to prevent over-acidification of
the thylakoid lumen. The high rates of CEF in hcef mutants has allowed more facile probing of
key CEF processes to explore its regulation and biochemical pathways. The genetic loci of some
of these hcef mutants have been identified (69, 79), yielding hints about the mechanism and
regulation of this process.
One high CEF plant (hcef1) was found with a mutation in the CBB cycle enzyme
fructose-1,6-bisphosphatase (69). Despite similar effects on overall photosynthesis, suppressing
the expression of glyceraldehyde phosphate dehydrogenase (70) and fructose bisphosphate
aldolase (79) led to increased CEF, while suppression of others enzymes, including Rubisco
small subunit (70), ATP synthase, and the Rieske protein of the b6f complex (80), or the triosephosphate/phosphate translocator (81), did not lead to an apparent increase in CEF. These
contrasting effects suggest that CEF, as discussed in more detail below, may be regulated by
specific intermediates or byproducts, rather than simply by slowing overall photosynthesis.
14

1.3.E.1 CEF pathways
The pathway(s) and regulatory mechanisms of CEF are currently the subject of debate.
The identity of the PQR is as yet unresolved and several protein complexes have been proposed,
but have yet to be fully characterized. Several putative PQRs, and many additional associated
proteins have been proposed (Figure 1.5), such as Proton Gradient Regulation 5 (PGR5) and the
PGR5-Like protein (PGRL1) (82, 83). Some of these are plant-specific, such as the NADPH
dehydrogenase complex (NDH) (84, 85) or algae-specific, such as type II (non-proton pumping)
NADH:plastoquinone oxidoreductase, NDA2 (86). It should be noted that these pathways are not
necessarily mutually exclusive, and may either operate in parallel (87–90) or differentially under
specific conditions (79, 91, 92).
1.3.E.1.A Chloroplast NDH pathway
One proposed route of CEF is through the thylakoid NAD(P)H dehydrogenase complex
(NDH) (84, 85). The NDH complex is highly conserved in cyanobacteria and green plants across
all phyla (93). The ~550 kDa (84) plant complex consists of >24 subunits, 11 of which are
chloroplast encoded (94) and are (with a few notable exceptions) highly homologous to
mitochondrial and bacterial respiratory Complex I (85). The identification of novel plant-specific
NDH-associated proteins suggests that the plastid NDH has diverged in structure and regulation
from Complex I and cyanobacterial NDH-1 (94, 95). Complex I contains an integral membrane
hydrophobic arm with at least 3 proton channels (85); the genes coding for these channels are
conserved in NDH (94, 96).
In Complex I, proton pumping is attributed to conformational changes in conserved
lysine and glutamate residues that are also conserved in chloroplast NDH (96–98). The specific
and generally high level of conservation of the NDH complex in vascular plants suggests that the

15

Figure 1.5 Proposed routes of cyclic electron flow around photosystem I. 1) The thylakoid
NAD(P)H dehydrogenase complex (NDH). 2) A PS I-b6f supercomplex. 3) A Type II (nonproton pumping) NAD(P)H dehydrogenase complex (NDA2). 4) A ferredoxin plastoquinone
reductase thought to be comprised of a complex containing PGRL1 and PGR5.

16

same mechanism operates in plant chloroplasts. The occurrence of proton pumping in NDH
would make CEF a very efficient ATP-generating process and imply an important role for the
NDH complex in maintaining the thylakoid ATP budget.
In support of involvement of NDH in CEF, the hcef1 mutant shows constitutively high
CEF that requires the NDH complex (but not PGR5, which is involved in an alternative pathway;
see below). These results suggest that NDH can catalyze high rates of CEF, at least under certain
conditions (69). Direct participation of NDH in photosynthesis is further supported by a report
that NDH is activated by formation of a supercomplex with PS I, implying a direct participation
in photosynthesis rather than a function in non-photosynthetic plastids (95).
Because chloroplast NDH levels are relatively low (NDH:PS I ~1:20) (95), it is not clear
whether this enzyme is able to catalyze high rates of CEF under all conditions. However, NDH
subunits accumulated at much higher levels under environmental stresses (99–101) and in hcef1
(69). Increased expression is also seen in C4 bundle-sheath chloroplasts, where the required ratio
of ATP/NADPH is higher (91). It has also been observed that CEF and NDH activity increase in
vitro and in vivo upon exposure to H2O2, which is expected to increase during environmental
stress (99, 102–104) (discussed further in Chapter 3). Thus, NDH may be activated, both by
increased expression and at the enzyme level, to catalyze CEF that is highly efficient in ATP
synthesis under environmental stresses. On the other hand, it is almost certain that other CEF
pathways operate in other systems and under different environmental conditions, as discussed in
the following sections.
1.3.E.1.B Supercomplex pathways
While Chlamydomonas reinhardtii lacks genes for chloroplast NDH (105), a CEF
competent supercomplex comprised of PS I, LHCI, LHCII, the b6f complex, ferredoxin NADP
17

+

reductase (FNR), and PGRL1 has been isolated (92). In C. reinhardtii, it has been observed that
CEF activation correlates with an antenna state transition to PS I (106). State transitions
modulate partitioning of photosynthetic antenna complexes between PS I and PS II (39). State 1
is induced when PS I is preferentially excited, resulting in the continued association of LHCII
with PS II, and under these conditions LEF is operating (107). When light conditions favor PS II,
the redox state of the PQ pool activates phosphorylation of LHCII by the STT7/STN7 kinase and
LHCII associates with PS I in state 2 (108). It is under these state 2 conditions that the
supercomplex was isolated (92). Similar supercomplex formation has been suggested to
facilitate CEF in higher plants (3, 109), but has not been fully characterized. In addition, NDH is
proposed to require formation of a supercomplex with PS I (95).
Overall, it appears that rearrangement of thylakoid protein complexes into supercomplexes associated with processes like state transitions may regulate CEF and other
photosynthetic functions. It may therefore be important that C. reinhardtii appears to have a
much larger capacity for state transitions than higher plants: up to 80% vs ~15% (39, 108),
respectively, consistent with the reported higher capacity for CEF in C. reinhardtii.
The specific electron-transfer pathway for supercomplex CEF is not known, although it
has been proposed that heme ci of the b6f complex may act as a conduit for electrons from the
stroma to the plastoquinone reductase (Qi) site (110). It is also intriguing that the CEF
supercomplex shows relatively high rates of electron transfer from the b6f complex to P700
despite the fact that the experiments were performed in isolated complexes with plastocyanin
concentrations much lower than those expected in thylakoids (92). The channeling of electron
transfer by close association in super-complexes could be important for controlling the fraction

18

of electrons sent into LEF or CEF, as discussed in Breyton et al. (111), Peng et al. (95), and
Johnson (65).
1.3.E.1.C The ferredoxin-plastoquinone reductase (FQR) pathways
CEF has also been proposed to be facilitated by an antimycin A-sensitive PQR termed
ferredoxin:quinone reductase (FQR) (112). While its identity has yet to be fully elucidated, this
complex has been proposed to be a complex comprised of PGRL1 and PGR5 (82, 83, 113).
Shikanai and coworkers (82, 87) identified a mutant (pgr5) lacking the ability to maintain
pmf-dependent NPQ and deficient in the small membrane-associated protein PGR5. This mutant
was determined to be lacking the ability to perform FQR-mediated PQ reduction, based on in
vitro reduction assay kinetics similar to antimycin A inhibition. Avenson et al. (35) calculated
that the PGR5 pathways could contribute up to 13% of proton flux in vivo under steady-state
conditions in A. thaliana. However, they also showed that many of the phenotypes associated
with pgr5 can be attributed, not to CEF, but to effects on ATP-synthase activity, calling into
question the physiological importance of PGR5-catalyzed CEF. Similarly, Aro and coworkers
(114) have presented evidence that the main function of PGR5 is in the regulation of PS I redox
state, perhaps via effects on ATP synthase, and PGR5 is not involved in CEF.
Work on the hcef1 mutant likewise implied that PGR5 is dispensable for CEF (69). The
hcef1 pgr5 double mutant retained the same elevated rates of CEF as hcef1, implying that, at
least in this high CEF mutant, PGR5 is not needed. In contrast, elevated CEF was completely
lost in the hcef1 crr2-2 double mutant that lacks NDH (69), implying that NDH is required for
elevated CEF, either directly as a PQ reductase or indirectly via regulatory or developmental
effects. Elevated CEF in tobacco plants with anti-sense-suppressed glyceraldehyde phosphate
dehydrogenase (gapR) showed no sensitivity to antimycin A, an inhibitor of the FQR pathway

19

(70). This is in agreement with the proposed role of PGR5 in regulation of LEF but not CEF
(114). Similar observations were made with another high CEF mutant, fba3-1 deficient in
fructose-1,6-bisphosphate aldolase (79). In this mutant, however, it has been proposed that the
PGR5-dependent pathway may compensate for the absence of NDH. Overall, these results do not
support PGR5 as the primary route of CEF, but rather suggest multiple roles for PGR5 in
maintaining photosynthetic processes (92).
In addition to its proposed role in the C. reinhardtii super-complex, PGRL1 has recently
been suggested to act as a PQR (113). PGRL1 is an integral thylakoid-membrane protein (83)
that associates with PGR5 and PS I in higher plants, and associates with the CEF supercomplex
in C. reinhardtii (92). Loss of PGRL1 leads to a decrease in Fd-dependent dark PQ reduction as
assayed by chlorophyll a fluorescence, similar to the loss seen in PGR5 (82). Identification of
conserved cysteine residues, binding of a metal cofactor, and the ability of recombinant PGRL1
to reduce dimethyl-p-benzoquinone (DMBQ) (dependent on the presence of PGR5) suggests that
PGRL1 is a quinone reductase (113). However, some caution with regard to this interpretation is
warranted. Inhibition of the observed FQR activity, performed with artificial quinones, required
antimycin A concentrations 3 orders of magnitude higher than are effective in thylakoids (82,
112, 115).
1.3.E.1.D NDA2 pathway
For C. reinhardtii, it has been proposed that CEF is catalyzed by a type II
NADH:plastoquinone oxidoreductase, termed NDA2, related to those found in bacteria, fungi,
plants, and protists (86). NADH, which is reduced during metabolism but not directly by PS I,
appears to be the preferred substrate for NDA2, suggesting a primary role in metabolism-related
PQ reduction (i.e., not directly related to the light-driven CEF) (86). Consistent with this role,

20

NDA2 appears to be important for redox poising of the plastoquinone pool (116) and hydrogen
production under sulfur deficiency (117), i.e., functionalities not directly associated with CEF.
However, NDA2 also shows partial (10%) activity with NADPH (86), perhaps allowing it to
operate in CEF, although a contribution to photosynthetic ATP production has yet to be
demonstrated.
1.3.E.2 Regulation of CEF
Several signals for CEF regulation have been proposed, including CBB cycle
intermediates, state transitions, ATP/ADP ratios, NADPH or Fd redox states, and/or ROS (68,
70, 99, 102, 103, 106, 109, 111). Given the numerous pathways that may be involved, it is likely
that multiple regulatory processes are employed, and each pathway may have different signals
for activation. This is thus an active area of research, albeit with little resolution to date. A few
important advances are discussed in the following.
Livingston et al. (70) assayed major photosynthetic metabolites from mutants affected in
photosynthesis with elevated (hcef1 and gapR) and unchanged (anti-sense Rubisco small subunit
tobacco mutant; ssuR) CEF, respectively. None of the metabolites measured (fructose-6phosphate, dihydroxyacetone phosphate, fructose bisphosphate, phosphoglyercerate, ribulose-5phosphate, glucose-5-phosphate, phosphoenolpyruvate) correlated with the activation of CEF,
indicating that these metabolites are unlikely to be simple regulators of CEF.
It has been proposed that CEF in C. reinhardtii is regulated by state transitions (discussed
above). The transition to state 2 has been shown to correlate with activation of CEF in algal
systems under specific conditions (92, 106). It has been shown, however, that state 2 is not
absolutely required for CEF in C. reinhardtii (118) and more recent work has shown that CEF

21

can proceed in the absence of state transitions, leading to suggestions that redox status, and not
phosphorylation state, controls CEF (119, 120).
Consistent with a role in balancing the chloroplast’s energy budget, it has been suggested
+

that ATP/ADP or NADPH/NADP ratios can regulate CEF (109). CEF has been proposed to be
activated during photosynthetic induction, thus increasing ATP supply when this supply is
limiting the CBB cycle (64, 109). However, this conclusion is not consistent with previous
spectroscopic findings (121). Furthermore, metabolic profiling of hcef mutants showed no
+

significant differences in ATP/ADP or NADPH/NADP ratios in wild type versus mutant lines
with elevated CEF (70), suggesting that regulation of CEF may instead involve ROS.
ROS have been implicated in a number of signaling processes, such as growth and
environmental stress response (122–124). The hcef1 mutant accumulates significantly higher
levels of H2O2 compared to wild-type A. thaliana (104) (Chapter 3). With the discovery of the
role of H2O2 in the activation of the NDH complex by phosphorylation (102), CEF was
established as a target of ROS. Indeed, infiltration with H2O2 also leads to increased CEF, the
level of which correlates with the concentration of H2O2 (104) (discussed in Chapter 3).
However, H2O2 has also been shown to inhibit several steps in the CBB cycle,
knockdown/knockout mutants of which show an hcef phenotype (70, 125). CEF in the hcef1 and
gapR mutants is independent of the antimycin A-sensitive pathway (69, 70). In hcef1, and
possibly also in gapR, ROS accumulation, or another signal downstream of CBB cycle
inactivation, may lead to phosphorylation and activation of NDH. Therefore, it is possible that
FQR or other paths of CEF are also regulated by additional factors independent of ROS.
22

1.4 Type II flexibility mechanisms: regulation of pmf partitioning and ATP synthase
activity and the consequences for NPQ
While Type I flexibility mechanisms alter the ATP/NADPH ratio available for
downstream metabolism, Type II mechanisms alter pmf without changing this ratio. Included in
the latter category is regulation of the ATP synthase and changes in pmf partitioning between
!pH and !". Downregulation of the ATP synthase increases pmf, and subsequently the qE
response, while alterations in pmf partitioning may alter qE response to total pmf.
1.4.A The chloroplast ATP synthase is a central regulator of photosynthesis
The enzymatic activity of the chloroplast ATP synthase is rapidly modulated in response
to metabolic or physiological conditions, allowing it to regulate both the light reactions, via
effects on the proton circuit, and the assimilatory reactions, via effects on chloroplast ATP and Pi
levels (25, 34). Under low CO2, environmental stresses, and feedback-limiting conditions at high
CO2, ATP synthase activity is rapidly and reversibly downregulated, slowing proton efflux from
the lumen (32, 34). The resulting buildup of pmf activates downregulation of the antennae via
processes underlying qE, and slows electron transfer at the b6f complex, thus downregulating the
light reactions (25).
Because many of the assimilatory enzymes of photosynthesis are controlled by stromal
ATP, ADP, and Pi levels, changes in the activity of the ATP synthase should act to co-regulate or
coordinate the light and assimilatory reactions (126). Decreasing ATP synthase activity should
result in decreased ATP and increased ADP and Pi, regulating assimilatory and metabolic
reactions.
23

Several possible mechanisms can be proposed for metabolism related ATP synthase
regulation. It has been well documented that chloroplast ATP synthase is regulated by redox
modulation of #-subunit thiols (127). Recently, Kohzuma et al. (128) further tested this
hypothesis by modifying the conserved acidic residues on the "-subunit. This manipulation
modified the redox potential of the thiols, but the mutant complex still maintained wild-type
responses to metabolic limitations. Thus, redox switching and metabolic regulation of ATP
synthases have different mechanisms and physiological roles. Redox regulation appears to act as
a sensitive on-off switch, activating ATP synthase even in very low light (129, 130), possibly to
prevent “wasteful” ATP hydrolysis in the dark (131).
In one possible model for metabolic regulation of ATP synthase, the availability of
stromal Pi can be drawn down to below its KM at the ATP synthase, e.g., under limiting internal
CO2 (32) or under feedback limitation resulting from accumulation of photosynthetic products
under high CO2 levels (132, 133). In agreement with these models, Takizawa et al. (126) found
that lowering Pi levels decreases the activity of ATP synthase, leading to downregulation of
photosynthesis. It has also been observed that downregulation of the ATP synthase occurs at
high CO2 levels, when Pi availability is thought to limit photosynthesis (134). These results are
consistent with, but do not establish, a direct causal relationship between Pi levels and ATP
synthase activity.
It has also been proposed that ATP synthase can be regulated by phosphorylation and
subsequent binding of a 14-3-3 protein (135, 136). Although the phosphorylation state of the
ATP synthase appears to change in response to light exposure of dark-adapted plants (137), no
measurements of phosphorylation in response to metabolic status have been reported. The 14-324

3-protein family is known to interact with enzymes involved in chloroplast metabolism (138),
but a direct role in phosphorylation of the ATP synthase has yet to be shown, and 14-3-3 proteins
may be nonspecific.
1.4.B Regulation of pmf partitioning
Chemiosmotic coupling can be driven by pmf via either of its two components, proton
diffusion potential, !pH, and transthylakoid electric field, !". Early work on isolated thylakoids
suggested that the contribution of !" to thylakoid pmf was low, but more recent work showed
that !" can make significant contributions to pmf (139). Although the two components of pmf
are thermodynamically equivalent, they have very different consequences for the regulation of
photosynthesis (25). Buildup of the !pH component decreases the pH of the thylakoid lumen,
activating qE and slowing electron transfer at the b6f complex (26). Modulation of the fraction of
!pH and !" to total pmf can change the sensitivity of qE to the pmf, and thus act as an
additional level of photoprotective regulation at limiting CO2, elevated temperatures, or other
environmental stresses (26, 33, 37, 140). While the mechanism of this modulation partitioning is
currently not understood, it is clear that several factors determine the storage of !", i.e.,
membrane capacitance, ionic strength, and buffering capacity of the lumen (139). It is unlikely
that thylakoid membrane capacitance would change sufficiently rapidly to account for the
observed changes in !"/!pH. On the other hand, ion channels in the plasma membrane,
chloroplast inner envelope and thylakoid membranes can control ion movements across the
thylakoid. In the absence of counter-ion movement across the membrane, we would expect the
buffering capacity of the lumen to maintain a higher fraction of !pH than !" (Figure 1.6A).
2+

2+

-

+

Active ion channels for Mg , Ca , Cl , and K have been confirmed to occur in thylakoids
25

Figure 1.6 Proposed mechanisms of changes in pmf partitioning.

26

Figure 1.6 (Cont’d) A) In the absence of counter ion movement, !" should predominate. B)
Counter ion movement (i.e. lumenal cation efflux/anion influx) would allow increased !pH. C)
A membrane permeable weak base would increase the buffering capacity of the lumen,
collapsing the !pH. Inspired by Nicholls and Ferguson (141).

27

(142–144), and Cruz et al. (139) proposed the regulation of chloroplast ion balance as the major
+

mechanism of adjusting !pH/!" (Figure 1.6B). In accord with this proposal, a gated K channel
was found to be required for maintaining !pH in cyanobacteria (145, 146). Consistent with the
view that pmf partitioning is controlled by ion movements, it was recently found that disruption
of a thylakoid K+ channel in plants resulted in higher partitioning of pmf into !", with
consequences for the regulation of photosynthesis by the proton circuit (147).
An additional regulatory mechanism was proposed by Ioannidis et al. (148) who showed
that putrescine, a polyamine acting as a weak base and involved in plant stress responses, can
regulate !"/!pH by modulating the effective proton buffering capacity of the lumen. This work
supports a “biological weak base” model (Figure 1.6C), in which molecules like polyamines may
allow for rapid modulation of pmf partitioning.
1.5 Concluding remarks
The proton circuit plays a central role in governing the responses of processes underlying
NPQ, predominantly by activating the qE response, but also by governing the rate of electron
transfer through the b6f complex and thus modulating the PQ-pool redox state. The proton circuit
also provides a very flexible means by, which metabolic or physiological states of the chloroplast
influence NPQ responses, e.g., via regulating CEF or ATP synthase. It is obvious that, although
we are beginning to delineate the interacting processes, a large number of important questions
remain open.
One way to assess whether our understanding of the proton circuit and its influence on
NPQ is complete, and to identify additional important open questions, is to compare
physiological responses with numerical simulations. There have been several attempts to model
28

pmf and the qE response to pmf (26, 139, 149). These models are based on broad simplifications,
but show qualitatively reasonable pmf and responses that reflect the general trends seen in vivo.
The recent model of Zaks et al. (149) is noteworthy in that it incorporates several possible
regulatory mechanisms for qE, and provides a solid framework for future inclusions of more
complex processes. However, these models also highlight the fact that we know little about
several regulatory mechanisms, especially CEF and pmf partitioning that are clearly important
for balancing the proton circuit (150).
1.6 Acknowledgements
We would like to thank Dr. Jeffrey Cruz and Dr. Nick Fisher for helpful discussions. The
authors are funded by Grant DE-FG02-11ER16220 from the Division of Chemical Sciences,
Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy.

29

Chapter 2
Redox regulation of the antimycin A sensitive pathway of cyclic electron flow around
photosystem I in higher plant thylakoids
Deserah D. Strand, Nicholas Fisher, Geoffry A. Davis, L. Ruby Carrillo, and David M. Kramer

30

2.1 Abstract
The chloroplast must rapidly regulate supply of reducing equivalents and ATP to
meet downstream metabolic demands. Cyclic electron flow around photosystem I (CEF) is
proposed to help balance the ATP/NADPH budget by using reducing equivalents to drive
plastoquinone reduction, leading to the generation of proton motive force and subsequent ATP
synthesis. While high rates of CEF have been observed in vivo, isolated thylakoids show only
very slow rates, suggesting that the activity of a key complex is lost or down-regulated upon
isolation. We performed a systematic investigation to discover the factors responsible for this
loss of activity. We show that isolation of thylakloids while in the continuous presence of thiol
reductant dithiothreitol (DTT) maintains high CEF activity through the ferredoxin-dependent
antimycin A-sensitive pathway. Maintaining low concentrations (~2 mM) of reduced DTT while
modulating the concentration of oxidized DTT leads to reversible activation/inactivation of CEF
with an apparent midpoint potential of –306 mV and n = 2, consistent with redox modulation of
a thiol/disulphide couple.
We propose the 2 most widely studied routes of CEF in plants, the FQR and NDH, are
both regulated by redox status of the chloroplast, but have different roles in energy balance. In
our model, FQR is activated under reducing conditions as a ‘first response’ and the NDH
activated later when the FQR fails to restore balance and ROS (and likely an oxidizing
conditions) is produced.
2.2 Introduction
The light reactions provide the ATP and reducing equivalents required for metabolic
processes within the chloroplast. Most photosynthetic energy is stored by the linear electron flow
(LEF) pathway, which involves activation of both photosystem II (PS II) and photosystem I (PS

31

I). Excitation of photosystem II (PS II) with photons results in the oxidation of water at the
oxygen evolving complex and the reduction of plastoquinone to plastoquinol (PQ/PQH2). PQH2
is oxidized at the Qo (quinol oxidation) site of the cytochrome b6f complex (b6f) in which one
electron is passed to plastocyanin in the lumen, and the other electron participates in the Q-cycle,
reducing a PQ bound at the Qi (plastoquinone reductase) site. Light energy is also captured by
the photosystem I (PS I) reaction center, resulting in oxidation of PC and reduction of ferredoxin
+

(Fd), which is used as a donor for downstream metabolism, including reduction of NADP to
NADPH. Proton deposition into the lumen is coupled to electron transfer reactions during water
oxidation and the Q-cycle, establishing an electrochemical gradient of protons, or proton motive
force (pmf) across the thylakoids. The pmf is, in turn, used by the ATP synthase to drive the
photophosphorylation of ADP to ATP. LEF in its unmodified form should provide a fixed
stoichiometry of reducing equivalents (i.e. reduced FD, NADPH) and ATP, therefore any
changes in the metabolic demands of the chloroplast must be met by rapid changes in alternative
electron flow to prevent formation of reactive intermediates (2, 3, 78). Multiple alternative
electron pathways have been identified including the water-water cycle (44, 45), the malate shunt
(41), the plastid terminal oxidase (59, 61, 151), and cyclic electron flow around photosystem I
(CEF) (112).
CEF can alleviate an ATP deficit by passing electrons from the acceptor side of PS I back
to PQ, driving the translocation of protons into the lumen without net reduction of NADP+. The
mechanism and role of CEF is intensely debated, partly because it is difficult to measure, and
partly because multiple proposed pathways may participate under different species and under
different conditions [see review in (2)]. Proposed pathways of CEF, in higher plants, include the
32

multi-subunit type I NADPH dehydrogenase complex (NDH) which is homologous to
respiratory Complex I (84, 85), the PGR5-PGRL1 dependent antimycin A-sensitive Fd
dependent quinone reductase (FQR) (82, 112, 113), and PQ reduction through the Qi site of the
b6f complex (110). In algae, CEF involves the formation of a b6f-PSI supercomplex (92), and/or
the FQR related PGR5 (152, 153) and PGRL1 (92, 154), or NDA2, a type II NADPH
dehydrogenase (86).
The regulation of CEF is also enigmatic, with multiple mechanisms proposed that may
impact different pathways , including ROS, state transitions, chloroplast redox state, ATP/ADP,
+

and NADPH/NADP (64, 70, 76, 92, 99, 102, 106, 111, 154, 155). Past work linking CEF
activation to antenna state transitions in Chlamydomonas has recently been challenged (120,
155).
A possible clue to the open questions surrounding CEF is that, while high rates of CEF
are readily measured in vivo (69, 70, 72, 79) (Chapters 3 and 4), this activity is severely
diminished in thylakoid preparations, suggesting that the a key component of the process is lost
or down-regulated upon isolation. We thus aimed in this work to systematically explore the
relationship between thylakoid in vitro conditions and CEF activity, revealing redox
requirements that may be involved in the regulation of CEF in vivo.
2.3 Results
2.3.A Effect of reductant on CEF activity in vitro
Freshly isolated thylakoids were assayed for their ability to generate a light driven proton
gradient (Figure 2.1). Assays were repeated under two conditions, one in the presence of methyl
viologen, allowing exclusively LEF, and the other in the presence of presence of NADPH and Fd

33

Figure 2.1 Light induced formation of pmf monitored by acridine fluorescence quenching
-2 -1

upon illumination with 200 µmol photons m s photosynthetically active radiation. A)
Linear electron flow, with methyl viologen (MV) as a final electron acceptor, black line, and
CEF, in the presence of DCMU, grey line, in chloroplasts isolated without any reductant B)
Linear electron flow, black line, and CEF, grey line, in chloroplasts isolated with 2 mM reduced
DTT C) CEF in chloroplast isolated with 2 mM reduced DTT, black line, and without DTT, grey
line D) Sensitivity of chloroplasts isolated with 2 mM reduced DTT, black line, to 5 µM
Antimycin A, grey line, and 20 µM nigericin, dashed grey line.
34

and dichloromethyl urea (DCMU), allowing CEF exclusively. Proton translocation activity was
determined by acridine fluorescence quenching under illumination, which indicates the
generation of the !pH component of pmf (156). This method of assessing CEF has the advantage
of being sensitive to true pmf-storing processes and not to recombination reactions within PS I
that can appear as CEF but do not result in net proton translocation. A 5:1 ratio of 9-amino
acridine (9AA) and 9-amino-6-6-chloro-2-methoxy-acridine (ACMA) was used to obtain a more
linear response of acridine fluorescence quenching to !pH (156). Valinomycin and KCl were
added to each reaction to dissipate the transthylakoid electric field component of pmf (!#) so
that all pmf was in the form of !pH, which is detectable by acridine fluorescence quenching.
In Figure 2.1A thylakoids were isolated without reductant in the isolation buffers. While
onset rate and extent of !pH generated in the presence of MV was substantial, that for CEF was
only 8.2% (estimated from the initial slope) that of LEF, indicating a low CEF rate of proton
translocation. The final steady-state amplitude of acridine quenching for CEF was only 3.8% that
of LEF. The relaxation of the !pH under the different conditions was similar, indicating that the
difference in !pH accumulation could not be attributed to changes in the proton efflux rates or
ATP synthase activities.
When chloroplasts were isolated in the presence of 2 mM reduced DTT, the relative
capacity for CEF increased substantially (Figure 2.1B) to 13.3% (initial slope) and 12.1%
(amplitude) of LEF. This increase translates to an increase in the initial rate of proton influx via
CEF of 63% when chloroplasts are isolated in the presence of reduced DTT and an increase in
the total amplitude of CEF generated CEF by 275% (Figure 2.1C). The similarity in slow
acridine fluorescence dark recovery kinetics across treatments indicates that the rate of proton

35

efflux was affected by the presence of DTT, and is consistent with a lack of substrate (ADP or
Pi) for the chloroplast ATP synthase.
CEF in the chloroplast preparation containing reduced DTT was sensitive to 5 µM
Antimycin A (75% decrease in initial slope, 64% decrease in the amplitude of quenching, Figure
2.1D), indicating involvement of the FQR (115), and was completely dissipated by the addition
of nigericin, which leads to complete uncoupling when used in the presence of valinomycin and
excess KCl (Figure 2.1D).
To confirm the activity of CEF we measured light-induced absorbance changes at 703 nm
+

to monitor the redox state of P700 (Figure 2.2). As oxidized P700 absorbs less at this
wavelength than reduced P700, in the presence of DCMU, the monitoring of 703 nm absorbance
changes allows the comparison of both the reduction state of the PQ pool, and rates of electron
transfer through PS I. Figure 2.2 shows light induced changes in !A 703nm in chloroplasts
isolated with 2 mM DTT and those isolated without. The light induced absorbance changes show
a slower oxidation of P700 in the chloroplasts isolated with 2 mM DTTRED than those without
(t1/2 = 2 s and t1/2 < 10 ms, respectively), indicating a more reduced PQ pool in the thylakoids
isolated with reductant. We also see a faster reduction of P700 in the dark interval following the
actinic flash in the presence of 2 mM DTTRED.
While all results were obtained with NADH, similar observations were seen with
NADPH (data not shown).
Interestingly, after isolation, 2 mM DTTRED was not able to restore full activity of CEF
in thylakoids isolated without reductant (Figure 2.3). Even when assayed with DTTRED, CEF
36

Figure 2.2 Light induced P700 redox changes. Chloroplasts were isolated and assayed with 2
mM reduced DTT, black line, and without reductant, grey line, in the presence of DCMU2.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
2
Data for this figure were kindly provided by Dr. Nicholas Fisher.
37

Figure 2.3 Irreversible loss of CEF in thylakoids isolated in the absence of DTT. Light
induced acridine fluorescence quenching in DCMU treated thylakoids isolated with 2 mM
DTTRED (black line) or without reductant (grey line). Activity was assayed in the presence of 2
mM DTTRED.

38

activity of the thylakoids isolated without DTTRED was only half that of the thylakoids isolated
with DTTRED (53.5% from initial slope, 57% from amplitude of acridine fluorescence
quenching), suggesting irreversible loss or damage to an FQR component.
2.3.B Dependency of Fd on CEF activity
The extent of formation of !pH in the presence of DCMU was sensitive to the addition of
Fd, with an 81% decrease in proton influx via CEF (initial slope) and a 71% decrease in total pmf
generated by CEF (amplitude) when Fd is omitted (Figure 2.4), indicating the DTTRED did not
reduce the PQ pool directly, but was dependent on Fd, further supporting involvement of the
FQR; these results are consistent with previous work (82, 112). In contrast, in ‘highly active
chloroplasts’ addition or absence of NADH had less effect on the extent of light driven acridine
fluorescence quenching in the presence of DCMU (34% decrease in proton influx and 19%
decrease in pmf, from initial slope and amplitude of fluorescence quenching, respectively)
(Figure 2.4).
As thylakoids were prepared from a crude preparation of broken and intact chloroplasts,
and ruptured within the reaction cuvette, it is likely there was a small amount of Fd and
+

NAD(P) /NAD(P)H remaining in the cuvette. This may explain residual activity without
addition of Fd.
2.3.C The redox dependence of CEF activation by DTT
In order to determine the redox potential in which CEF is active or inactive, we assayed
activity in a range of DTTOX/2DTTRED concentrations in thylakoids isolated with 2 mM DTT to
maintain activity (see above, Figure 2.3). The concentration of DTTRED was held constant at

39

Figure 2.4 Dependence of CEF on Fd in vitro. Acridine fluorescence quenching as an indicator
of pmf formation in the presence of DCMU. Total extent of pmf was greatly diminished without
addition of Fd, but was not largely changed when NADH was omitted from the reaction.

40

4 mM and incremental addition of DTTOX was added to obtain Figure 2.5. Figure 2.5 shows the
fractional activity of CEF determined by the total amplitude of acridine fluorescence quenching
as a function of the ratio of the [DTTOX/2DTTRED]. Half maximal CEF activity was observe at a
calculated redox potential of -306 mV versus standard hydrogen electrode when the data was fit
to a modified form of the Nernst equation. The fit is consistent with a slope of n = 2.
We tested this reversibility by rupturing ‘highly active’ chloroplasts in a concentration of
DTTOX/2DTTRED predicted to poise the redox potential at -300 mV, which should be near half
activity. This was followed by subsequent addition of 9 mM DTTRED, which should bring the
redox potential down to -380 mV, and strongly activate or restore CEF activity. Activity of CEF
was increased after addition of DTTRED (Figure 2.6).
This does not address whether CEF is reversibly lost during preparation as we have seen
previously (Figure 2.3), but shows that the regulatory component is rapidly reversible in vitro.
2.4 Discussion
There is a multitude of research conducted on CEF in vitro using isolated thylakoids from
mutants, or under varying conditions. A reduction in already low rates of CEF in thylakoids in
vitro have been used to infer large physiological effects in vivo (82, 87, 157). We have
demonstrated that the nature of isolation and assay conditions are vital for the conservation of
CEF in vitro.
2.4.A FQR-mediated CEF is active under reducing conditions in vitro
It has been previously described in algae that the redox state is the determining factor for
formation of a CEF competent supercomplex (92, 155). However, in plants, this is the first
demonstration that CEF is dependent on chloroplast redox state in vitro. While activation of the
41

Figure 2.5 Redox titration of CEF activity under varying concentrations of
DTTOX/2DTTRED. The Em of CEF activation is calculated to be around -306 mV.

42

Figure 2.6 Rapid reversibility of CEF. ‘Highly active’ chloroplasts were assayed for CEF
activity in partially oxidized DTT (black line), and then assayed for CEF activity after
subsequent addition of 9 mM DTTRED (grey line).

43

NDH complex has been suggested to be activated in the presence of ROS (99, 102) (discussed in
Chapter 3), we have demonstrated that the ‘elusive’ antimycin A sensitive FQR, possibly
involving a PGR5/PGRL1 complex, is more active under reducing conditions (Figure 2.1D, and
2.4). This lends support to two separate and independently regulated CEF pathways.
2.4.B FQR-mediated CEF may involve a regulatory thiol
The activity of CEF in isolated chloroplasts is dependent on the redox potential of the
reaction buffer (Figures 2.1A-C, 2.2, and 2.6) and the midpoint potential of activity is -306 mV
when fit with n = 2 (Figure 2.5). This range is in agreement with involvement of a thiol, which
have been described as having midpoint potentials ranging from as high as -124 mV to -330 mV
and may be shifted further with pH (158–160).
Assignment of CEF activity in our assays to the FQR is supported by both the sensitivity
of activity to antimycin A (Figure 2.1D) and the dependency of Fd (Figure 2.4). Based on this
identification of the FQR pathway, if a thiol is involved, it is possible the thiol is directly located
on the FQR related PGRL1. Keeping in mind the effect of pH on redox potential, this would be
in tentative agreement with recent work (113), in which a reversible thiol/disulfide pair (Em = ~280 mV) was identified within PGRL1, which may be involved in formation and disassociation
of the PGRL1-PGR5 complex.
There is extensive precedence for regulatory thiols in photosynthesis controlling both the
electron transport chain and carbon fixation reactions in response to chloroplast redox state. The
observation that CEF is also regulated in response to chloroplast redox state adds an additional
level of regulation to the proton circuit of photosynthesis.
Among the regulatory thiols associated with the electron transport chain is one with a Em
~-360 mV associated with activity of the b6f complex (161). It is possible the regulation of this
44

thiol could interfere with our measurements of in vitro CEF, however, as activity of the b6f
complex is apparently downregulated with a more reducing redox potential, the effect would
cause our apparent CEF activity to be underestimated. Another point of redox regulation of the
proton circuit is at the ATP synthase (131, 162). Again, with any interference at this regulatory
point, we would expect an underestimation of CEF in our active state or overestimation of CEF
in our inactive state.
The observation that FQR-mediated CEF is more active in a reduced state is in agreement
with the involvement of a regulatory thioredoxin recently described by Courteille et al. (163), but
the exact interactions are unclear as this enzyme has been proposed to be a CEF suppressor.
2.4. C FQR-mediated CEF is sensitive to thylakoid isolation conditions
The large increase in CEF when thylakoids are isolated with DTTRED is not seen when
DTTRED is added to thylakoids isolated without reductant. This suggests the irreversible loss or
damage of an FQR component. This lability of a complex has been seen in multiple instances of
redox active proteins such as ascorbate peroxidase, which is irreversible damaged when isolated
without ascorbate (164). This is important to note as rates of CEF may be underestimated if
complex activity is partially lost during isolation.
2.4. D Implication for regulation in vivo, two pathways of CEF
While addition of reduced DTT is not physiological, the conditions in the above
experiments may be reflective of conditions in which changes in downstream metabolism leads
+

to a more reduced NAD(P) /NAD(P)H pool and lead to rapid reduction of a regulatory thiol and
in turn, rapid activation of CEF (Figures 2.6 and 2.7A). This rapid activation would allow for a
much faster response to redox state than that shown for activation of NDH in response to H2O2
45

Figure 2.7 A comprehensive model of CEF activation in vivo. A) An ATP deficit leads to the
buildup of reducing power in the stroma which leads to the activation of the antimycin A
sensitive FQR pathway of CEF B) If FQR mediated CEF is unable to provide enough ATP to
supply downstream metabolism, the acceptor side of PS I becomes closed and electron transfer
to O2 results in ROS formation and activation of the NDH complex.

46

(99, 102) (Chapter 3). This more rapidly activation of CEF may be a ‘first response’ to balance
the ATP/NADPH output of the light reactions before the buildup of reducing equivalents leads to
ROS production. This ‘first response’ by the FQR would also likely have a lower ATP output
than the NDH complex, which may be capable of proton pumping. This may be beneficial to
rapidly modulate CEF with a lower ATP output when the reduction state of the chloroplast
indicates an imbalance forming. If this route of CEF is unable to augment the ATP deficit, the
formation of ROS may then signal for a slightly slower, but more efficient ATP producing route
of CEF, the NDH complex (Figure 2.7B), (discussed in Chapter 5).
2.5 Methods
2.5.A Chloroplast isolation
A crude mixture of broken and intact chloroplasts was isolated from market fresh
spinach, purchased the morning of experiments, using a protocol modified from (165). Leaves
were kept in the dark at 4°C until use. Leaves were ground in buffer (350 mM sorbitol, 50 mM
HEPES, 5 mM MgCL2, 2.5 mM EDTA, 0.1% w/v bovine serum albumin, pH 7.6), at a
concentration of 1g/mL, 3 times for 3 seconds each in a standard kitchen blender, then filtered
through 3 layers cheesecloth and 1 layer Miracloth. The filtrate was centrifuged for 10 minutes at
3000 X g. The supernatant was poured off and the pellet was resuspended in a small volume of
buffer (350 mM sorbitol, 50 mM HEPES, 5 mM MgCL2, 2.5 mM EDTA, pH 7.6) using a soft
bristle paintbrush and used immediately. All steps were carried out on ice. Reductant was added
to all isolation buffers at concentrations described in the results during assays, unless otherwise
noted. While extents of activity varied from each isolation, the experimental effects remained
constant. Therefore, we present representative results and comparisons are made between
chloroplast isolated simultaneously.
47

2.5.B Oxidation and reduction of dithiothreitol (DTT)
To obtain oxidized DTT, a solution at concentration was left at room temperature for a
week. The solution was monitored for oxidation by the appearance of the 283 nm peak in the
absorbance spectrum indicating oxidation (data not shown).
DTT was reduced by addition of 400 mM borohydride to a 200 mM DTT solution on ice
for 1 hour with occasional gentle swirling. Then HCl was added to a 400 mM concentration to
convert the borohydride to boric acid. This was allowed to incubate on ice for 30 minutes. The
solution was brought to pH 7.6 and diluted to 100 mM DTT concentration. Reduction of DTT
was confirmed by the disappearance of the 283 nm peak in the absorbance spectrum (data not
shown) indicating essentially complete reduction.
2.5.C Spectroscopic measurements
For all spectroscopic measurements, chloroplasts were ruptured in the reaction cuvette
prior to the experiment in low osmotic assay buffer (50 mM HEPES, 5 mM MgCl2, pH 7.6). For
LEF measurements the reaction buffer contained 50 $M methyl viologen as a final electron
acceptor. CEF measurements contained 20 $M DCMU, 25 $M ferredoxin, and excess NADH
(100 $M). Spectroscopic measurements were performed on an in-house built spectrophotometer
as described in Hall et al. (166) modified with a fluorimeter cuvette chamber. For the redox
titration, NADH was decreased to 25 µM to minimize the impact on redox potential.
2.5.C.1 Acridine fluorescence quenching
To monitor the accumulation of the light induced changes in the pH component of pmf in
thylakoids, acridine (80 $M 5:1 9-aminoacridine:ACMA) (156) was excited at 430 nm and total
fluorescence emission between 495 nm and 700 nm was detected by filtering with a FGL495

48

longpass filter and a BG18 shortpass (Schott) filter (ThorLabs). Fluorescence quenching was
measured during a dark-light-dark interval, and normalized to the initial dark fluorescence level.
2.5.C.2 Absorbance changes at 703 nm
The redox state of PS I was measured using absorbance changes at 703 nm as described
in (120) using a spectrophotometer modified from Hall et al. (166). Contribution of chlorophyll
a fluorescence was limited by addition of 1 mM hydroxylamine followed by a 2 second actinic
flash. Any residual chlorophyll a fluorescence signal contamination was monitored by repeating
the experiment with a reference wavelength (730 nm) and determined to be minimal under these
conditions.
2.6 Author contributions
All data presented, except where noted, were obtained by Deserah D. Strand. However,
significant intellectual contribution was also made by the co-authors. Geoffry A. Davis and L.
Ruby Carrillo supplied a large amount of preliminary data in collaboration, and therefore have
been added as co-authors to this manuscript. L. Ruby Carrillo and Dr. Nicholas Fisher tested
many different chloroplast isolation protocols and buffers. Geoffry A. Davis and Dr. Nicholas
Fisher contributed significantly to the optimization of the acridine fluorescence quenching
protocols and concentrations of reaction buffer components.
2.7 Acknowledgements
We would like to thank Dr. Jeffrey Cruz for helpful discussions. The authors are funded
by Grant DE-FG02-11ER16220 from the Division of Chemical Sciences, Geosciences, and
Biosciences, Office of Basic Energy Sciences of the US Department of Energy.

49

Chapter 3
Activation of cyclic electron flow around photosystem I by hydrogen peroxide in vivo3
Deserah D. Strand, Aaron K. Livingston, Mio Satoh-Cruz, John E. Froehlich,
Veronica G. Maurino, and David M. Kramer

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
3
Preliminary data for this manuscript has previously appeared in Livingston (104) and this work
is referenced within.
50

3.1 Abstract
Cyclic electron flow (CEF) around photosystem I is thought to balance the ATP/NADPH
energy budget of photosynthesis, requiring that its rate be finely regulated. The mechanisms of
this regulation are not well understood. We observed that mutants that exhibited constitutively
high rates of CEF also showed elevated production of H2O2. We thus tested the hypothesis that
CEF is activated by H2O2 in vivo. CEF was strongly increased by H2O2 both by infiltration or in
situ production by chloroplast-localized glycolate oxidase, implying that H2O2 can activate CEF
either directly by redox modulation of key enzymes, or indirectly by affecting other
photosynthetic enzymes. CEF appeared rapidly, within 20 min of exposure to H2O2, suggesting
activation of existing CEF complexes. H2O2-dependent CEF was not sensitive to antimycin A or
loss of PGR5, indicating that increased CEF probably does not involve the PGR5-PGRL1
associated pathway. In contrast, the rise in CEF was greatly suppressed in a mutant deficient in
the chloroplast NADPH:PQ reductase (NDH), supporting the involvement of this complex in
CEF activated by H2O2. We propose that H2O2 is a missing link between environmental stress,
metabolism, and redox regulation of CEF in higher plants.
3.2 Introduction
In oxygenic photosynthesis, linear electron flow (LEF) is the process by which light
energy is captured to drive the extraction of electrons and protons from water and transfer them
through a system of electron carriers to reduce NADPH. LEF is coupled to proton translocation
into the thylakoid lumen, generating an electrochemical gradient of protons !!!!!! !!or proton
motive force (pmf). The pmf drives the synthesis of ATP to power the reactions of the Calvin51

Benson-Bassham (CBB) cycle and other essential metabolic processes in the chloroplast. The
pmf is also a key regulator of photosynthesis, in that it activates the photoprotective qE response
to dissipate excess light energy, and down-regulates electron transfer by controlling the rate of
oxidation of plastoquinol at the cytochrome b6f complex (b6f), thus preventing the buildup of
reduced intermediates (1, 3).
LEF by itself results in the transfer or deposition into the lumen of three protons for each
electron transferred through PS II, plastoquinone (PQ), the b6f complex, plastocyanin and
photosystem I (PS I) to ferredoxin (Fd). The synthesis of one ATP is thought to require the
passage of 4.67 protons through the ATP synthase, so that LEF should produce a ratio of
ATP/NADPH of about 1.3; this ratio is too low to sustain CBB cycle or supply ATP required for
translation, protein synthesis transport or other processes (2). The relative demands for ATP and
NADPH should dynamically change depending on environmental, developmental and other
factors, so that the regulation of ATP/NADPH must be dynamically regulated.
Several alternative electron flow pathways in the chloroplast have been proposed to
augment ATP production thus balancing the ATP/NADPH budget of the chloroplast (2, 3).
Perhaps the least understood of these pathways is cyclic electron flow around photosystem I
(CEF), in which electron flow from the acceptor side of PS I is shunted back into the PQ pool,
generating additional pmf that can power ATP production with no net NADPH production. There
are several proposed CEF pathways that may operate under different conditions or in different
species (2, 3). In higher plant chloroplasts, the most studied routes of CEF are (A) the antimycin
A sensitive pathway, which involves a complex of two CEF-related proteins, PGR5 and PGRL1,
directly reducing the quinone pool (82, 87, 112, 113), (B) the respiratory Complex I analog, the

52

NADPH dehydrogenase (NDH) complex (84, 85, 94) which oxidizes Fd or NADH to reduce
plastoquinone (85, 167), and (C) through the Qi site of the b6f complex (110, 168). Different CEF
mechanisms appear to operate in other species. In Chlamydomonas, for example, CEF appears to
be conducted by a supercomplex of PS I, the b6f complex, and the PGRL1 protein (92, 155) and
the involvement of PGR5 has recently been described as important for CEF under hypoxia (153).
Regardless of the mechanism of CEF, the overall process must be precisely regulated to
properly balance the production of ATP to match the demands of metabolism. The mechanism of
this regulation is not known, but many general models have been proposed. Perhaps the most
widely cited regulatory model is the antenna state transition, which were previously shown to be
correlated with activation of CEF in C. reinhardtii (92, 106) and favor the formation of the PS Ib6f supercomplex (92). However, it was recently shown that state transitions are not required for
CEF activation, supporting models that include redox control (118, 120, 152, 153, 155) [see also
(169)]. Other possible regulatory mechanisms include sensing of ATP/ADP ratios (64, 109), the
redox status of NAD(P)H or Fd (111), various CBB metabolic intermediates [reviewed in (70)],
calcium signaling (102, 154), phosphorylation of CEF complexes (102) and the reactive oxygen
species (ROS) H2O2 (70, 99, 102, 103).
In this work, we concentrate on the possible role of H2O2 which is produced by the light
reactions of photosynthesis and are already known to regulate other cellular processes such as
plant growth, development, and defense (170–172). Based on in vitro studies, it was previously
proposed that H2O2 could activate CEF or chlororespiration by modifying the NDH complex
(102). It has also been shown that H2O2 can increase the expression of the NDH complex (99)
53

and may further affect the accumulation of photosynthetic metabolites, indirectly activating CEF
(70). Consistent with this possibility, H2O2 is a well-documented signaling molecule (173),
possibly through its ability to oxidize thiols (174, 175). Furthermore, H2O2 is expected to be
produced under many conditions that initiate CEF, e.g. under a deficit of ATP, when electrons
should accumulate in the PS I acceptor pools leading to superoxide production which can be
converted to H2O2 by superoxide dismutase (45).
This study aims to test the hypothesis that CEF can be initiated in vivo by H2O2 using a
combination of in vivo spectroscopy and genetic modifications to selectively and rapidly initiate
H2O2 production in the chloroplast.
3.3 Results
3.3.A H2O2 accumulation in the high cyclic mutant hcef1
The Arabidopsis mutant, hcef1 (69), which is deficient in chloroplastic fructose-1,6bisphosphase (FBPase) activity, and displays constitutively high CEF rates, was checked for
increased H2O2 accumulation. When H2O2 concentration is measured by resorufin fluorescence,
the product of the reaction between Amplex Red and H2O2, hcef1 has 3 times as much peroxide
as the Col-0 (Figure 3.1, 2.99 ± 0.17 and 1.00 ± 0.13, respectively, n = 3, p > 0.001, student’s ttest). Additionally, when stained with 3,3’ diaminobenzidine (DAB), leaves of hcef1 (Figure
3.1, inset B) showed qualitatively increased levels of H2O2 when compared to the wildtype
strain, Columbia-0 (Col-0) (Figure 3.1, inset A). These results indicate that the loss of FBPase
led to increased H2O2 accumulation, probably by a buildup of reducing intermediates of
54

Figure 3.1 Relative H2O2 accumulation in hcef1. Quantitatively determined H2O2 content of
leaves based on resorufin fluorescence against a standard curve and normalized to the average
Col-0 H2O2 content. Data is based on a per area basis. Mean ± SD, n = 34. Inset: Qualitative
H2O2 accumulation measured by 3,3’ diaminobenzidine (DAB) staining of representative leaves
of Col-0 (A) and hcef1 (B)5

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
4
This data was kindly provided by Dr. Mio Satoh-Cruz.
5
This data appears previously in (104) and has been generously provided by Dr. Aaron
Livingston.!
55

photosynthetic electron transfer, and are consistent with a correlation between H2O2 and CEF
activation. We thus set out to determine if this relationship could be causal.
3.3.B Effects of H2O2 production by plants expressing glycolate oxidase in chloroplast
We next tested if CEF is activated in the presence of H2O2 by comparing photosynthetic
properties of Col-0 and transgenic Arabidopsis plants that express glycolate oxidase (GO)
targeted to the chloroplast (176). These “GO” plants produce H2O2 in the chloroplast by the
oxidation of glycolate upon activation of photorespiration, and are thus useful tools for studying
changes in photosynthetic activities induced by the metabolic generation of H2O2 in the
chloroplast. We focus mainly on the GO5 line because of its relatively high and robust H2O2
production rates (176, 177) (Figure 3.2), but we obtained similar results with other lines
(discussed below).
GO expression had measurable effects on several photosynthetic parameters (Figure 3.3).
PS II photochemical efficiency ($II) (Figure 3.3A) and LEF (Figure 3.3B) saturated more rapidly
with increasing light intensity in GO5 compared to Col-0, leading to a lower LEF, particularly at
-2 -1

intensities above 200 µmol photons m s . GO5 showed stronger activation of qE, response
(Figure 3.3C), likely indicating an increase in light-induced pmf and lumen acidification. A
substantial (about 2-fold) increase in pmf in GO5 was confirmed by the extent of the darkinterval electrochromic shift (ECS) absorbance changes, ECSt (Figure 3.3D) (40–42, see
Materials and Methods). This increased pmf occurred in GO5 despite lower LEF, suggesting that
either proton influx was increased above that supported by LEF or that proton efflux was slowed.

56

Figure 3.2 Relative H2O2 accumulation in GO5.!Quantitatively determined H2O2 content of
leaves based on resorufin fluorescence against a standard curve and normalized to the average
Col-0 H2O2 content. Data is based on a per unit chlorophyll basis. Mean ± SD, n = 3. Assays
were performed on leaves from fully mature rosettes just prior to bolting as described for
spectroscopic assays.

57

Figure 3.3 Effects of chloroplast targeted glycolate oxidase expression on photosynthesis.

58

Figure 3.3 (Cont’d) Comparison of photosynthetic parameters of Col-0 (black circles) and GO5
(open circles). Panels A-E show, as a function of light intensity; Photochemical efficiency of
photosystem II ($II , Panel A); Linear electron flow (LEF, Panel B); rapidly reversible exciton
quenching (qE, Panel C); relative light-induced pmf, as measured by the total decay of the ECS
+

(ECSt, Panel D) ;and proton conductivity (gH ) of the ATP synthase, as estimated by the
relaxation kinetics of the ECS signal (panel E). Panel F shows ECSt as a function of the
estimated pmf generated from LEF alone (pmfLEF). Mean ± SD (n = 3).

59

+

The conductivity of the thylakoid to protons (gH ), which primarily reflects the activity of the
+

ATP synthase, was estimated by the decay kinetics of the ECS signal (34, 178). In GO5, gH was
approximately 30% lower than in Col-0 (Figure 3.3E), implying that, although the ATP synthase
activity was somewhat decreased in the mutant, it could not by itself explain the large increase in
light-induced pmf, suggesting an increase in CEF [see discussion in (33, 69)]. This conclusion
was supported by a statistically significant (ANCOVA p < 0.05, n = 3) increase of about 36% in
light-driven pmf, estimated by ECSt as a function of pmfLEF (Figure 3.3F), a parameter that
estimates pmf generated by LEF alone (33, 179). These results indicate an increase in pmf above
that attributable to LEF alone, suggesting that CEF was activated in GO5.
Increased CEF in GO5 was independently confirmed by comparing estimated light+

driven proton flux (vH ) as a function of LEF (26, 33). As shown in Figure 3.4A, the slope of vH
as a function of LEF was increased in GO5 in comparison to Col-0 by approximately 47.6%
(Figure 3.4A, ANCOVA p < 0.05, n = 3). The increase in CEF was eliminated by infiltration of
100 µM methyl viologen, which blocks CEF by shunting electrons from PS I to O2 (26) [see
+

Figure 4.3A in (104)]. This increase in ratio of CEF/LEF (!vH / vH

+

LEF,

see Eq. 3.3, Materials

and Methods) was not caused simply by decreasing LEF. Instead, assuming a similar ratio of
proton translocation for electron flux, CEF increased in absolute terms from minimal activation
-2 -1

in the control [as previously reported in (69)] to approximately 24 µmol electrons m s when
-2 -1

LEF was 50 µmol electrons m s .!
Analyzing the ECS and fluorescence data described in Eq. 3.3 (Materials and Methods),
+

we estimated that vH was increased by about 50% in GO5 over Col-0. The difference in the
60

+

-GO5

-WT

-NDH-18

+

Figure 3.4. Increased CEF in GO5. A) light-driven proton flux (vH ) vs LEF in attached Col-0
(squares) and GO5 (circles) leaves B) Calculated relative CEF from intact leaves and leaves
infiltrated with 20 µM antimycin A (AA). Mean ± (A) SD (n = 3) or (B) SEM (n % 3). *
Indicates statistical significance from Col-0, no significant differences were seen between water
and AA within genotype. B, inset) NDH-18 levels in GO5 grown at ambient CO2. Lanes were
loaded with 10 µg total chlorophyll6.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
6
Figure 3.4B, inset was kindly provided by Dr. John Froehlich.
61

apparent extents of CEF activation (about 36% versus about 50%) may be ascribed to differences
in proton to electron stoichiometries for CEF versus LEF or to ambiguities in quantification of
the proton and electron fluxes. Nevertheless, the multiple estimates of increased CEF were
qualitatively consistent, and correlated well with the extent of H2O2 production in multiple GO
lines with differing levels of GO activity (104, 176) [see Figure 4.3A-C in (104)]. In another
independent assessment of CEF induction, we observed a strong increase in GO5 compared to
Col-0 of the postillumination chlorophyll fluorescence rise (Figure 3.5), which is attributed to
CEF-related reduction of the plastoquinone pool in the dark through the NDH complex (79, 84,
180).
Together, these results strongly imply that introduction of GO in chloroplasts induces
increased CEF. However, in the GO plants, glyoxylate produced by the glycolate oxidase in the
chloroplasts, also accumulates, and may induce CEF. To test this hypothesis we measured CEF
in GO5 plants also coexpressing malate synthase targeted to the chloroplast (GOMS1 and
GOMS14 lines), which can further convert glyoxylate to malate (176). We observed qualitatively
similar photosynthetic effects of GO5 and GOMS14 (Figure 3.6), indicating that the production
of H2O2 rather than glyoxylate is the likely inducer of CEF.
The observed increase in CEF was insensitive to infiltration with 20 µM antimycin A
(Figure 3.4B, ANCOVA, p > 0.05, n = 3), well above the observed Ki for inhibition of the
antimycin A sensitive pathway of CEF (115). There is also an increase in NDH content of GO5
when grown at ambient CO2 levels (Figure 3.4B, inset). These results are consistent with the
post-illumination rise (Figure 3.5) in suggesting the involvement of NDH in GO5 elevated CEF.

62

Relative Fluorecence

GO5

Col-0

20s

Figure 3.5 Increased dark reduction of the PQ/PQH2 pool in GO5. Transient chlorophyll
-2 -1

fluorescence rise in the dark after illumination with 150 µmol photons m s (635 nm). Col-0
(black) and GO5 (red) representative data of 3 independent experiments. The full data set is
normalized to FM, while the inset data is normalized to [0,1] for comparison.

63

Figure 3.6 Increased CEF in mutants co-transformed with glycolate oxidase (GO) and
+

malate synthase (MS) targeted to the chloroplast. Relative CEF calculated from vH /LEF (Eq.
3.3) in Col-0, GOMS1 and GOMS14. Mean ± SD (n = 3).

64

3.3.C Kinetics of CEF induction upon activation of H2O2 production in GO5
Figure 3.7. shows the kinetics of induction of CEF in the GO5 plants upon rapid initiation
of H2O2 production. GO5 plants were initially grown under high (3000 ppm) CO2 conditions to
minimize photorespiration (181, 182), therefore preventing the production of H2O2 by GO (176,
177). Photosynthetic parameters were measured in intact leaves under 2000 ppm CO2 and then
rapidly switched to ambient air (about 400 ppm CO2) at time 0, activating H2O2 production.
Steady state fluorescence and ECS measurements were made every 14 min and analyzed as in
Figure 3.3A to estimate changes in CEF. Increased CEF appeared with a half time of about 21
+

min, reaching an apparent maximum relative CEF of 0.62, or a 62% increase in vH / LEF, in 42
+

min (!vH / vH

+

LEF,

see Eq. 3.3, Materials and Methods). No increase in CEF was seen in

similarly treated Col-0 leaves (Figure 3.7).
3.3.D Induction of CEF by infiltration of leaves with H2O2
In addition to using the GO plants to study CEF, preliminary studies showed a similar
response with infiltration of Col-0 with exogenous H2O2 [Figure 3.2 in (104)]. Infiltration of
leaves with as low as 300 µM (0.001%) H2O2 led to induction of significant CEF rates in vivo
[Figure 3.2 in (104)]. The observed H2O2-induced increase in CEF depended on the
concentration of H2O2 in the infiltrate, with an apparent half-saturation concentration of about
0.01%. Infiltration of Col-0 with 0.001%, 0.01% and 0.1% H2O2 increased relative CEF by 21%,
42% and 68% over water infiltrated leaves, respectively. In addition, infiltration of the mutant
65

Figure 3.7 Induction kinetics of H2O2 activated CEF. CEF activation timecourse in Col-0
(closed circles) and GO5 (open circles). Relative CEF as a function time after switch from 2000
ppm CO2 to 400 ppm CO2. Reported values are the mean of 3 biological replicates ± SEM.

66

pgr5 (82), lacking the antimycin A sensitive-PGR5 dependent CEF pathway, resulted in
activation of CEF at near Col-0 levels (65%), while the NDH deficient mutant, crr2-2 (94, 183),
accumulated very low levels of CEF (5%) [Figure 3.7 in (104)].
In addition, we assayed leaves incubated in 0.1% H2O2 for an increase in the
postillumination chlorophyll fluorescence rise as seen in the GO5 mutant (Figure 3.6). Electrons
in the stroma are used to detoxify H2O2, and it likely we are consuming these rapidly in the
presence of H2O2. In the steady state there is a continuous supply of electrons being fed into the
system via PS II to fuel CEF plastoquinone reductions, while in the dark our supply cannot be
replenished. In order to get a reproducible response to H2O2 infiltration, we modified the
experiment to include two far-red flashes during the dark interval and performed the
measurements under nitrogen. This allows the system to be poised and supply substrate to our
plastoquinone reductase. After infiltration with 0.1% H2O2 we noticed higher F0 values, as well
as higher FS values than in the water control (Figure 3.8), which we interpret as an already more
reduced quinone pool. When the light is shut off, the decay does not reach water levels. The
initial decay was followed with a short far-red pulse to oxidize the quinone pool. After the farred is turned off, we see a faster and overall larger increase in fluorescence, indicative of rereduction of the quinone pool in the peroxide treated leaves (Figure 3.8, inset). These results
suggest an increase in the rise via dark quinone reduction, and are consistent with steady state
measurements of increased CEF activity in response to peroxide infiltration (104).

67

Figure 3.8 Increased dark reduction of the PQ/PQH2 pool in leaves infiltrated with 0.1%
H2O2 for 3 hours. Transient chlorophyll fluorescence rise in the dark after illumination with
-2 -1

150 µmol photons m s (635 nm) under 100% N2 gas. Water (black) and 0.1% H2O2 (red)
representative data of 2 independent experiments.

68

3.4 Discussion
3.4.A CEF correlates with, and is induced by, H2O2 production in vivo
Past work has shown that mutants in higher plants that accumulate highly reducing
stromal redox components also induce elevated CEF, e.g. mutants deficient in FBPase (i.e.
hcef1), aldolase, and glyceraldehyde-3-phosphate dehydrogenase (69, 70, 79). Similarly, certain
environmental conditions, e.g. drought stress, lead to both increased H2O2 and elevated CEF (72,
184). In contrast, simply decreasing the rate of photosynthesis without increased of H2O2, does
not appear to induce CEF (50, 70).
These observations suggest a regulatory link between H2O2 production and the activation
of CEF. To test this possibility, we used transgenic GO plants, which express a chloroplast
targeted glycolate oxidase, and conditionally produce H2O2 under photorespiratory conditions.
Activation of H2O2 production in GO5 had strong effects on photosynthesis, decreasing LEF,
increasing thylakoid pmf and activating qE (Figure 3.2). Most strikingly, elevated H2O2
production led to strong activation of CEF (Figures 3.4 and 3.3F). CEF was similarly induced by
infiltration of leaves with H2O2 [Figure 3.7, (104)], suggesting that the effects were primarily
caused by H2O2 and not by GO-induced changes in metabolic intermediates. The activation of
CEF was likely too rapid (halftime of about 20 min, Figure 3.7), to involve de novo protein
synthesis, which is expected to be considerably slower (185, 186), suggesting that H2O2 can
activate pre-existing CEF machinery. This doesn’t exclude long term activation by accumulation
of CEF enzymes as seen in previous studies (99) and Figure 3.4B (inset), but instead suggests an
69

additional mechanism of rapid induction of CEF to meet fluctuating ATP demands as proposed
by Lascano et al. (102).
3.4.B H2O2 activates the antimycin A insensitive pathway
H2O2-induced CEF was insensitive to antimycin A (Figure 3.4B), implying that it does
not involve the PGR5/PGRL1 pathway (82, 112, 113), suggesting instead the involvement of the
chloroplast NDH complex, as previously shown in hcef1 (69). In support of this possibility, an
increase in NDH content was seen in GO5 (Figure 2B, inset) and the!activation of CEF by
infiltration with H2O2 was inhibited in the NDH-deficient crr2-2 mutant (104), while the mutant
deficient in PGR5 was activated to Col-0 levels. In addition, the NDH complex has been
associated with increased activity in the presence of oxidative stress (99, 102), and its activation
results in increased post-illumination fluorescence rise, which was increased in the GO5 mutant
and after infiltration with H2O2 (Figures 3.5 and 3.8).
3.4.C A regulatory role for H2O2 in activation of CEF?
We emphasize that CEF is likely to be regulated at a number of processes in different
species and under different conditions. Recent work by Takahashi et al. (155) and Lucker et al.
(120) suggest that CEF in Chlamydomonas is regulated by stromal redox status, but it is not yet
clear which redox components are involved. It is important to point out, though, that CEF in
higher plants and Chlamydomonas likely involve distinct protein complexes, mechanisms and
regulatory processes (86, 92).
Based on these results, we propose the hypothetical model (Figure 3.9) in which H2O2
production can regulate the activation of CEF. A deficit of ATP in the stroma should prevent the

70

Figure 3.9 Model of CEF activation in response to H2O2. A) Imbalances in downstream
metabolic processes lead to an excess of reductant in the stroma. B) This leads to increased
oxygen reduction from photosystem I and generation of superoxide anion. C) Superoxide is
dismutated to H2O2 and this signals the phosphorylation of NDH, activating CEF to supply ATP
for metabolism, without net reduction of NADPH.
71

turnover of assimilatory reactions, and cause a buildup of reductants (NADPH, Fd and PS I
acceptors) leading to generation of superoxide and H2O2. When the rate of H2O2 production
exceeds that of its detoxification by the water-water cycle (45), it will accumulate and interact
with one or more proteins regulating the activity of the antimycin A insensitive pathway of CEF,
possibly via a signal cascade leading to the phosphorylation of NDH as suggested by Casano et
al. and Lascano et al. (99, 102), or by inactivating CBB enzymes leading to secondary redox or
metabolic signaling.
3.5 Methods
3.5A Plant material and growth conditions
All plants, Col-0, GO plants (expressing glycolate oxidase targeted to the chloroplast,
37), and hcef1 (69) were grown in soil under growth chamber conditions with 16 h light of white
-2 -1

light (~80 µmol photons m s ), 8 h dark photoperiod, and a 22°C/18°C (day/night) cycle.
Where noted, plants were grown under the same conditions but at high (3000 ppm) CO2 to
prevent the production of H2O2 and GO through the photorespiratory pathway.
3.5.B In vivo spectroscopic assays
Under ambient CO2 conditions, the GO plants present with a slightly smaller, pale, and
patchy leaf phenotype, however, the chlorotic phenotype disappears at maturity. Therefore, all
spectroscopic measurements were made using intact fully expanded leaves in 25-30 day old
plants just prior to bolting. To fully induce differential H2O2 production, GO5 and all controls
-2 -1

were pre-illuminated for 1 hour at 350 µmol photon m s . Plants were then subsequently darkadapted for 10 min prior to analysis. Actinic light intensities ranged between 50-600 µmol

72

-2 -1

photons m s . Chlorophyll a fluorescence yield changes and light induced absorbance changes
were measured using on a spectrophotometer/fluorimeter (166) using the techniques described in
(69). Saturation pulse chlorophyll a fluorescence yield parameters (F0, FM, FS, FM’, FM’’,) were
recorded as described in (7, 32, 33, 179), using 1 s saturation pulses of approximately 10,000
-2 -1

µmol photons m s . These measurements were used to estimated the yield of PS II ($II), LEF,
and the rapidly reversible component of non-photochemical quenching, qE (187, 188). LEF was
calculated as:

Eq 3.1
!!! ! ! ! !!!

Where i is the actinic light intensity. Leaf absorptivity of the GO plants did not differ
significantly from Col-0 (p = 0.78, n = 3).
The ECS measurements were normalized for variations in leaf thickness and
pigmentation by the extent of the rapid rise single-turnover flash induced ECS (33, 69). The
ECSt and tECS parameters were taken from a first-order exponential decay fit as described in
(179). The pmfLEF parameter, estimating relative extents of pmf attributable to LEF, was
calculated as:

Eq 3.2
!"#!"# ! !"#!!! ! !!

73

Postillumination transient chlorophyll fluorescence transients were measured as described
in (43).
3.5.C Infiltration of leaves with antimycin A and H2O2
Infiltrations of 20 µM antimycin A and 0.1% H2O2 were carried out soaking detached
leaves between 2 saturated lab tissues in darkness for 3 hours. Successful infiltration with
antimycin A was confirmed by loss of the transient fluorescence quenching during
photosynthetic induction as previously described (189).
3.5.D Manipulation of gas concentrations
Humidified gas mixtures and ambient air were supplied to the underside of the leaf unless
indicated otherwise. Fluctuating CO2 concentrations were obtained using a gas mixer (LI-COR
6400) connected to a CO2 gas cylinder. N2 gas was supplied by a tank of 100% N2 (Airgas).
3.5.E Measurement of H2O2 production in leaves
Hydrogen peroxide was detected by DAB staining as described in (104), and by resorufin
fluorescence. Resorufin is the product of the peroxidase catalyzed reaction with Amplex Red and
H2O2. H2O2 was extracted from leaf discs in 50 mM phosphate buffer (pH 7.6) and incubated
with 2U horseradish peroxidase and 10 µM Amplex Red (Invitrogen) and resorufin fluorescence
was normalized to the average Col-0 H2O2 content. For GO5 plants, leaf age was as describe
above for spectroscopic assays. For hcef1, leaf age was as described for spectroscopic
measurements in Livingston et al. (69).

74

3.5.F Protein extraction and western blot
Total leaf protein was extracted from fully mature rosettes (described above) as described
in (69) and normalized to chlorophyll. Western blotting was performed as described in (69),
using anti-NDH-18 (T. Shikanai, Kyoto University).
3.5.G Quantifications of CEF
To quantify relative CEF, which should be proportional to the increase in slope seen in
Figure 3.3A, we used an approach modified from (104) and (120), in which relative CEF is
+

+

expressed as a fractional increase over the vH due to LEF (! vH / vH

+

LEF)

and is calculated by:

Eq. 3.3
!"#$%&'"!!"# ! !! !!!"#!!"#! ! ! !!!"#! !!!!"#!

+

where !!!"#!!"#! and !!!"#! !are the linear slope of vH plotted against LEF in treated or mutant
+ -

leaves and untreated or Col-0 leaves respectively. If the H /e of CEF was known, this could
potentially be converted to electron flux, however, in higher plants possessing NDH, it is
+ -

possible that CEF involves a proton pump that could significantly increase H /e (discussed in
chapter 5). Therefore, we have chosen to omit this conversion and present relative CEF as the
+

fraction increase of vH over LEF.
3.5.H Statistical analysis
Descriptive statistics and figures were generated using Origin 9.0 software (Microcal
Software), and statistical analyses were performed using MATLAB R2012a (The Mathworks) or

75

Microsoft Excel. All p-values less than 0.05 were considered statistically significant and are
noted within the text.
3.6 Author contributions
All data presented, except where noted, were obtained by Deserah D. Strand. However,
significant intellectual contribution was also made by the co-authors. Specifically, Veronica G.
Maurino suggested the experimental setup in Figure 3.7. and Aaron K. Livingston generated
extensive preliminary data, which has been referenced within, as well as a preliminary draft of
the manuscript.
3.7 Acknowledgements
The authors would like to thank Drs. Nicholas Fisher, Atsuko Kanazawa, and Jeffrey
Cruz for helpful discussions. We would like to also thank Dr. Toshiharu Shikanai for his kind
donation of pgr5 and crr2-2 seed. Work performed at MSU was funded by Grant DE-FG0211ER16220 from the Photosynthetic Systems program from Division of Chemical Sciences,
Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy
(to David M. Kramer). The development of the GO mutants was funded by the Deutsche
Forschungsgemeinschaft (DFG) through grant MA2379/11-1 (to Veronica G. Maurino).

76

Chapter 4
Uncoordinated expression of chloroplast proteins leads to H2O2 accumulation and activation of
cyclic electron flow around photosystem I7
Deserah D. Strand, Aaron K. Livingston, Mio Satoh-Cruz, Tyson Koepke, Heather M. Enlow,
Jeffrey A. Cruz, Deepika Minhas, Kim K. Hixson, Kaori Kohzuma, Amit Dhingra, Mary Lipton,
and David M. Kramer

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
7
Preliminary data for this manuscript has previously appeared in (104).
77

4.1 Abstract
In recent work, we identified a class of mutants in Arabidopsis that exhibit high rates of
cyclic electron flow around photosystem I (CEF). Here we describe a new member of this class,
high cyclic electron flow 2 (hcef2), possessing a nonsense mutation in the TADA1 (tRNA
adenosine deaminase arginine) locus, coding for a plastid-targeted tRNA editing enzyme
required for proper codon recognition. Both hcef2 and tada1 show increased thylakoid proton
motive force, light-driven proton translocation relative to electron fluxes and increased dark
reduction of plastoquinone, indicating increased CEF activity. Similar results were obtained for
mutants defective in chloroplast translation, including prsp3, which is partially deficient in
protein translation at the level of chloroplast ribosome. These mutants showed elevated H2O2,
which we have previously shown to increase CEF. Inhibitor sensitivity and protein expression
levels imply that increased CEF in hcef2 occurs through the NADPH:plastoquinone
oxidoreductate (NDH) pathway. Intriguingly, high-resolution proteomics showed that hcef2
accumulated abnormal stoichiometries of proteins of multi-subunit complexes, including
photosystems I and II, the cytochrome b6f complex, and ATP synthase. These findings suggest
that loss of coordination of plastid protein levels leads to accumulation of misassembled
complexes, ROS production and subsequent activation of CEF through the NDH complex.
4.2 Introduction
In higher plants, light is harvested by two distinct photochemical reaction centers,
photosystem II (PS II) and photosystem I (PS I) that stimulate electron transfer through series of
redox carriers to store solar energy in forms to drive biochemical processes (3). When PS I and
PS II are electronically connected in series, they drive linear electron flow (LEF), which results
+

in the oxidation of water and the reduction of NADP to NADPH. The electron transfer reactions
78

of LEF are coupled to the uptake of protons from the chloroplast stroma and their deposition into
the lumen, establishing an electrochemical gradient of protons, or proton motive force (pmf).
Protons are taken up during reduction of plastoquinone at the QB site of PS II and the Qi site of
the cytochrome b6f complex (b6f). Protons are released into the lumen during water oxidation at
the oxygen evolving complex (OEC) of PS II and during plastoquinol oxidation at the Qo site of
the b6f complex. The pmf generated in these electron and proton transfer reactions drives the
synthesis of ATP at the chloroplast ATP synthase.
The pmf is also a central regulator of the light reactions through its effects on lumen pH
dependent components of non-photochemical quenching (NPQ) (1), and electron flow through
the b6f complex (26, 190). The pmf, in turn, is modulated in response to environmental and
metabolic conditions (32, 34, 72, 78) (Chapter 1), coordination of photosynthetic electron
transport and carbon metabolism. The major points of pmf regulation lie at the ATP synthase and
with cyclic electron flow (2, 25, 78). Both of these processes have different effects on the
stoichiometry of the photosynthetic products ATP and NADPH produced from the light
reactions. While modulation of the ATP synthase results in unaltered ATP:NADPH output of the
light reactions, CEF increases the ATP:NADPH output.
Cyclic electron flow around PS I (CEF) involves PS I but not PS II (112). Electrons from
PS I are transferred to plastoquinone (PQ) forming plastoquinol (PQH2), which is subsequently
oxidized by the b6f complex and shuttled back to PS I by plastocyanin. The uptake of protons in
the Q-cycle, catalyzed by the b6f complex, results in translocation of protons from the chloroplast
stroma to the lumen and thus contributes to the formation of pmf and ATP synthesis without net
79

reduction of NADPH (115). In this way, CEF can act to augment an ATP deficit, and has thus
been implicated in balancing the chloroplast energy budget, and is thus thought to be
physiologically important under conditions of elevated ATP demands. For example, CEF is
induced under environmental stresses including drought (72, 191), high light (157), and chilling
(192) where ATP may be needed to repair cellular machinery, maintain ion homeostasis,
transport proteins, etc. CEF is also thought to supply ATP for CO2 concentrating mechanisms,
including the C4 cycle in plants (91) and the carbon concentrating mechanism (CCM) in green
algae (120), and appears to be critical under anoxia (153) or when CO2 is limiting (120), in the
green alga Chlamydomonas reinhardtii.
It is also possible that CEF plays a regulatory role in photosynthesis by acidifying the
thylakoid lumen and thus activating the photoprotective qE response and slowing electron flow at
the b6f complex (82, 157). It is important to recognize that uncontrolled activation of CEF will
also result in a change to the ATP/NADPH output stoichiometry, a situation that can lead to
deleterious secondary effects. Thus, chloroplasts also have alternate mechanisms of regulating
lumen acidification that do not result in alteration of ATP/NADPH, including modulation of
ATP synthase rates, and these appear to play primary roles in regulating photoprotection
(reviewed in Chapter 1).
Several alternative CEF pathways have been proposed, that involve different PQ
reductases, including the PGR5/PGRL1 dependent, antimycin A sensitive pathway (82, 87, 112,
153), the Qi site of the b6f complex (110), and the NAD(P)H dehydrogenase complex (NDH)
(84, 85). It is possible that different CEF pathways are activated in different species, and/or

80

under different conditions. To make matters more complex, a range of regulatory signals have
been proposed for CEF, including sensing of ATP/ADP ratios (64, 109), chloroplast redox status
(111, 119, 152, 153), metabolic intermediates (68), state transitions (92, 106), calcium (154), and
reactive oxygen species (99, 102, 104) (see Chapter 3).
Previously, we initiated an effort to discover new CEF structural and regulatory
components by isolating mutants of Arabidopsis (Arabidopsis thaliana) with constitutively
elevated CEF, which we named high cyclic electron flow (hcef) mutants (69, 70). The first of
these mutants to be reported, hcef1, was mapped to a missense mutation in the chloroplasttargeted fructose 1,6 bisphosphatase (FBPase), and appears to indirectly activate CEF by
disrupting redox balance (70) (see Chapter 3), possibly by activating a futile metabolic cycle that
consumes ATP (69). Results from a series of double mutants and inhibitors indicate that CEF
activated in hcef1 involves the chloroplast NDH complex and not the proposed PGR5/PGRL1
pathway.
In this work, we report on the isolation and characterization of hcef2, which was mapped
to an unexpected locus involving tRNA editing. Despite a very different mutation, and proteomic
consequences, hcef2 is found to have similar levels of CEF activation and photoprotection as
hcef1. This has strong implications for the role of CEF, including the possible involvement of
reactive oxygen species in its regulation, and the critical importance of strict regulation of plastid
proteome stoichiometries.
4.3 Results
4.3.A Genetic selection of hcef mutants
As described in Livingston et al. (69),we selected hcef mutants using a multi-stage
selection process from a pool of ethyl methanesulfonate (EMS) mutagenized seeds (Columbia

81

ecotype [Col-0], Lehle seeds [M2E-02-05]). We first screened for plants that displayed high
photoprotective ‘exciton quenching’ (qE) phenotypes using chlorophyll a fluorescence imaging.
We then subjected this population to secondary screening using direct measurements of lightdriven electron transfer, based on analysis of chlorophyll a fluorescence and proton fluxes based
on analysis of the electrochromic shift (ECS) signal (7, 69, 178, 179) to identify mutants with
elevated CEF.
4.3.B Growth of hcef2
The hcef2 mutant grew photoautotrophically in soil, at a decreased rate as described in
(104) and shown in Figure 4.1. The hcef2 mutant displayed a slightly pale appearance
owing to lower accumulation of chlorophyll compared to Col-0 levels per leaf area (157.8
2

2

mg/m ± 5.7 and 271.6 mg/m ± 5.1 respectively, p = 0.00001, student’s t-test, n = 3).
4.3.C Photosynthetic electron transfer properties of hcef2 compared to Col-0: responses of
the photosynthetic electron transport to the hcef2 mutation
Chlorophyll a fluorescence was used to estimate linear electron flow (LEF) and qE
responses, accounting for differences in leaf absorptivity using the approach described in (69,
193). The hcef2 mutant showed suppressed LEF rates across all light intensities used, about 4-

-2 -1

fold lower than Col-0 at saturating light (21.2 ± 1.7 and 82.6 ± 2.8 $mol e m s , respectively,
-2 -1

~480 $mol photons m s ) (Figure 4.2A). The half saturation irradiance for LEF in hcef2 (~90
-2 -1

-2 -1

$mol photons m s ) was ~65% that of Col-0 (~140 $mol photons m s ). In addition to
differences in PS II electron transfer rates, Col-0 and hcef2 have distinct 77K fluorescence
emission spectra (Figure 4.2B), reflecting large changes in the composition and state distribution

82

hcef2

Col-0
1 cm

Figure 4.1 Growth phenotype of hcef2 at 24 days. Plants were grown photoautotrophically on
-2 -1

soil with a 16:8 light/dark photoperiod at ~100 µmols photons m s , white light. For
interpretation of the references to color in this and all other figures, the reader is referred to the
electronic version of this dissertation.

83

Figure 4.2 Effects of hcef2 on photosynthetic properties. A) Photosynthetic linear electron
flow (LEF) as a function of light intensity. Col-0 (closed circles) and hcef2 (open circles). Mean
± SD, n = 3. B) 77k emission spectra. Col-0 (solid line) and hcef2 (dashed line). Emission
spectra are representative of 3 independent experiments. C) Energy dependent exciton quenching
(qE) as a function of light intensity. Col-0 (closed circles) and hcef2 (open circles). Mean ± SD, n
= 3. D) Photoinhibition (qI) as a function of light intensity. Col-0 (closed circles) and hcef2 (open
circles). Mean ± SD, n = 3.

84

of photosynthetic antenna complexes. The hcef2 mutant shows a large increase in the relative
emission of long-wavelength (~735 nm) emission associated with PS I antenna complexes,
compared to shorter wavelength (685 nm) emission reflecting antenna complexes associated with
PS II [for review see (194)]. In addition, the PS I associated peak showed a strong blue shift,
likely reflecting dissociation or loss of LCHI complexes from the PS I core (see below).
Col-0 showed typical sigmoidal responses of qE to light intensity (Figure 4.2C), with an
-2 -1

apparent half-saturation point at 375 µmol photons m s reaching approximately 0.9 at the
highest light intensity tested, similar to previous results on plants grown under similar conditions
(69). The photoprotective responses of hcef2 were distinct from those of Col-0, exhibiting high
-2 -1

levels of qE even at low light intensities. For example, at 80 µmol photons m s , qE in hcef2
was ~4-fold higher than in Col-0 (0.54 ± 0.026, 0.09 ± 0.12, respectively) (Figure 4.2C). In
-2 -1

hcef2, qE reached a maximum at about 280 µmol photons m s with a value of 0.73 (± 0.055, n
= 3), and decreased slightly at higher intensities. This decrease in qE likely reflects the
accumulation of photodamage at higher light in hcef2, which is seen as an increase in
photoinhibition, or qI (Figure 4.2D). Estimates of qI in hcef2 are likely even lower than reported
due to an already low maximal photochemical efficiency of PS II in dark-adapted leaves of hcef2
(0.59 ± 0.066) compared to Col-0 (0.80 ± 0.003).
4.3.D Responses of the photosynthetic proton circuit of hcef2
We analyzed the dark interval relaxation kinetics of the electrochromic shift (ECS) to
analyze the proton circuit of photosynthesis. The extent of light-driven pmf was estimated from
+

the total amplitude of the decay signal (ECSt); the relative rate of light-driven proton flux (vH )

85

was estimated from the initial slope of the ECS decay; and the conductivity of the thylakoid
+

membrane to protons (gH ), which reflects the activity of the chloroplast ATP synthase, was
estimated from the lifetime of the ECS decay (78, 178, 179). From these values we calculated
relative electron and proton fluxes through thylakoid components, and inferred the activation
state of CEF (69, 70).
The hcef2 mutant showed strongly decreased LEF compared to Col-0 (Figure 4.2A), yet
produced substantially higher light-driven pmf, as indicated by increased ECSt values as a
-2 -1

function of LEF (Figure 4.3A). At an LEF value of 20 $mol electrons m s , hcef2 had a 4-fold
higher ECSt than Col-0 (2 ± 0.1 and 4.2 ± 0.075 miliabsorbance units, respectively, n = 3). The
increased pmf is associated with qualitatively elevated qE in hcef2 (Figure 4.3B), as would be
expected based on the lumen pH-dependence of the qE response (1). While Col-0 showed a
sigmoidal dependence of qE on ECSt, as previously reported (26), hcef2 showed high activation
of qE at even low ECSt values, but saturated at relatively low LEF or ECSt extents (Figures 4.3B
and 4.3C). The higher sensitivity of qE responses in hcef2 was more sensitive to estimated pmf
changes (ECSt) in hcef2 compared to Col-0, indicating additional factors beyond the pmf play a
role in modulating the qE response in hcef2 (discussed below).
As discussed earlier (32, 78), thylakoid pmf can be increased with respect to LEF by
either accelerating proton influx through CEF or retarding proton efflux from the lumen by
inactivating the chloroplast ATP synthase. To distinguish between these possibilities, we
+

assessed the relative proton conductivity of the thylakoid membrane (gH ) using the ECS decay
86

Figure 4.3 Effects of hcef2 on the photosynthetic proton circuit. A) Light driven
transthylakoid pmf as measured by ECSt as a function of LEF. Col-0 (closed circles) and hcef2
(open circles). Mean ± SD, n = 3. B) qE as a function of LEF. Col-0 (closed circles) and hcef2
(open circles). Mean ± SD, n = 3. C) qE as a function of ECSt. Col-0 (closed circles) and hcef2
+

(open circles). Mean ± SD, n = 3. D) Thylakoid proton conductivity (gH ) as a function of light
intensity. Col-0 (closed circles) and hcef2 (open circles). Mean ± SD, n = 3.

87

+

lifetime measurements. As shown in Figure 4.3D, gH values for Col-0 and hcef2 were nearly
identical, varying by less than 10%, indicating that the observed increases in pmf and qE
responses in hcef2 could not be explained by down-regulation of the chloroplast ATP synthase.
4.3.E Estimates of CEF in hcef2
We next used three different approaches to directly assess the activation of CEF in hcef2.
In the first approach, we compared proton flux estimated from initial decay rates of the ECS
+

+

signal (vH ) with LEF estimated from chlorophyll a fluorescence parameters. When vH was
plotted as a function of LEF (Figure 4.4A), hcef2 showed a ~3-fold increase in slope over Col-0
(0.0039 ± 0.0011 and 0.0013 ± 0.0003, respectively, p = 0.0103 ANCOVA, n = 3), indicating an
+ -

increase in the light-driven fluxes of protons over LEF. Because LEF produces a fixed H /e

stoichiometry, the additional protons would need to be supplied independently of PS II, i.e. by
activation of CEF.
In the second approach (Figure 4.4B), we compared relative light-driven pmf, estimated
by the ECSt parameter with calculations of the pmf from LEF alone (pmfLEF) (35). This approach
is based on different assumptions and largely independent of extrinsic factors, such as the leaf
content of ECS-responding carotenoids, etc. (33, 179). The dependence of ECSt on pmfLEF was
approximately 3-fold higher in hcef2 compared to Col-0 (0.0037 ± 0.0005 and 0.0012 ± 0.0003,
respectively, p < 0.0001 ANCOVA n = 3), indicating that hcef2 accumulates larger extents of
pmf than can be attributed to changes in LEF, supporting the conclusion that CEF is strongly
activated in hcef2. It should be noted that estimates of LEF by analysis of chlorophyll
fluorescence depend on the fraction of light energy absorbed by PS II. The 77K fluorescence

88

Figure 4.4 Activation state of CEF in Col-0 and hcef2.
+

A) Light driven transthylakoid proton flux (vH ) as a function of LEF. Col-0 (closed circles) and
hcef2 (open circles). Mean ± SD, n = 3. B) ECSt as a function of pmf generated by LEF (pmfLEF).
Col-0 (closed circles) and hcef2 (open circles). Mean ± SD, n = 3.

89

emission spectra (Figure 4.2B) show a decrease in the relative fluorescence of PS II at 685 nm
relative to that attributable to PS I at about 735 nm, possibly indicating a decrease in PS II
relative to PS I excitation. In any case, the possible error introduced by this antenna change
would lead to an overestimate of LEF, and therefore an underestimation increased CEF for the
data in Figures 4.4A and 4.4B.
In the third approach we measured postillumination changes in chlorophyll a
fluorescence that indicate the non-photochemical reduction of the PQ pool associated with
activation of the NDH-pathway for CEF (79, 84, 85, 180). Typically, such fluorescence rise
experiments are conducted by exposing leaves to continuous illumination for a few minutes and
the fluorescence yield is followed after switching off the light. An initial decrease of
fluorescence is caused by rapid oxidation (on tens to hundreds of milliseconds time scale) of QA
by PQ. When NDH is active, this initial phase is followed by a slower fluorescence rise as PQ
becomes progressively reduced by NDH. During initial trials, we found that the decay and rise
phases were more clearly resolved when leaves from growth conditions were partially darkadapted (for 10 min) and exposed to short (10 ms duration) pulses of intense actinic light. As
shown in Figure 4.5, each pulse resulted in increased fluorescence yield reflecting light-induced
reduction of QA. The fluorescence yield then deceased in multiple phases after each flash. A
rapid phase, with a half time of less than a few ms, reflected the equilibration of QA and PQ
redox states in the dark. In Col-0 (Figure 4.5A, black line), each pulse resulted in progressively
more reduced PQ pool as indicated by the increases in dark fluorescence levels. This
interpretation was confirmed by the decreased in fluorescence yield induced by far-red (730 nm)
illumination (Figure 4.5A, gray line), which preferentially excites PS I photochemistry resulting

90

Figure 4.5 Kinetics of postillumination fluorescence rises in Col-0 and hcef28.
Relative fluorescence yield changes in Col-0 (A) and hcef2 (B) in the dark after a series of
actinic flashes (black) and changes with far-red illumination after the actinic flashes (gray). Data
is representative of 3 independent experiments.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
8
Data for Figure 4.5 was kindly provided by Dr. Jeffrey Cruz.
91

in net oxidation of the PQ pool and QA. Cessation of far-red illumination resulted in a slow
return to higher fluorescence yields indicating the activity of a PQ reductase. These phenomena
were also observed in hcef2 (Figure 4.5B, black line), but were stronger and more rapid. In fact, a
distinct fluorescence rise phase was seen after the third flash in hcef2 that we interpret as
indicating strong activation of PQ reductase activity. The interpretation was confirmed by
application of far-red illumination (Figure 4.5B, gray line), which resulted in substantial
quenching of the signal. Rise occurred after each additional flash and continued during the
following dark period. We conclude that hcef2 has a substantially higher activity of PQ reductase
than Col-0.
4.3.F Antimycin A infiltration of hcef2
+

In Col-0 we observed no significant differences in the ratio of vH /LEF between leaves
infiltrated with water or 20 µM antimycin A (Figure 4.6, 0.0015 ± 0.0025 and 0.00158 ±
+

0.00026, respectively p > 0.05 n = 3). The elevated ratio of vH /LEF in hcef2 was also unaffected
by 20µM antimycin A (Figure 4.6, 0.00551 ± 0.0014 and 0.00461 ± 0.00095, respectively p >
0.05 n = 3), indicating hcef2 CEF is antimycin A insensitive.
4.3.G Identification of the genetic locus of hcef2 as TADA1
Map based cloning and deep sequencing was used to identify the probable genetic locus
for the hcef2 mutation to a point mutation in TADA1 (At1G68720). This C>T mutation
introduces a stop codon at R643 (Figure 4.7), eliminating the C-terminus of the protein
containing the active site required for function (195, 196).
The T-DNA insert line GK-119G08 contains an insertion in the first exon in At1G68720
(196). Similar to hcef2, tada1 showed strongly increased CEF as indicated by a 5-fold increase
92

+

Figure 4.6 Antimycin A insensitivity of CEF in hcef2. Slope of vH /LEF in leaves infiltrated
with either water or antimycin A (AA). Mean ± SD, n = 3.

Figure 4.7 Introduction of a stop codon into the TADA1 locus in hcef2. Translation is
terminated before the active site of TADA1, leading to the loss of function9.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
9
The genetic mapping leading to the creation of hcef2 leading to the creation of Figure 4.7 was
performed by Dr. Mio Satoh-Cruz.
93

+

relative to Col-0 of vH /LEF (0.0063 ± 0.003 and 0.0011 ± 0.0004, respectively, p < 0.05 n = 3)
(Figure 4.8A) as well as in ECSt/pmfLEF (0.0057 ± 0.001 and 0.0011 ± 0.0003, respectively, p <
0.001, n = 3) (Figure 4.8B) relationships. These results indicate increased CEF equal to or greater
than hcef2 and support the identification of hcef2 mutation within TADA1. Likewise, GK119G08 showed no statistical difference from hcef2 in leaf chlorophyll content or absorptivity
(data not shown).
Delannoy et al. (196) showed that the tada1 phenotype was reversed by expressing the Cterminus of TADA1 behind a 35S promoter. We transferred this construct by crossing the
complimented tada1 mutant with hcef2 followed by segregation and genotyping for
homozygocity of the hcef2 mutation, lack of the tada1 insertion, and possession of the
P35S:!NTADA1 construct. Verified lines were analyzed spectroscopically for suppression of
+

the hcef2 phenotype. The increased vH as a function of LEF seen in hcef2 (Figure 4.4A) was
completely suppressed in the hcef2 P35S:!NTADA1 line (Figure 4.9), i.e the slope returned to
Col-0 values (0.0013 ± .00029 and 0.0013 ± 0.00031, respectively, p > 0.05, ANCOVA n = 3).
These results confirm TADA1 as the site of the mutation causing elevated CEF in hcef2.
4.3.H Translational defects in the chloroplast lead to increases in CEF.
The TADA1 gene codes for a tRNA editing enzyme, suggesting that a defect in
translation machinery somehow leads to increased CEF. To test if this effect is a general
consequence of decreased chloroplast translation efficiency, we assayed for increased CEF in
mutants defective in nuclear encoded peripheral ribosomal proteins. The prsp3 mutant contains a
T-DNA insert in the At1g68590 locus with a complete loss of PRSP3 (197). The rps17 mutant
contains a T-DNA insert in the At1g79850 locus, resulting in decreased expression of RPS17 by
94

+

Figure 4.8 Elevated CEF in tada1. The ratios of A) vH /LEF and B) ECSt/pmfLEF of intact
leaves were measured and estimated as described in Material and Methods and Figure 4.4. Date
represent the mean ± SD with n = 3. Asterisks indicate statistical difference from Col-0.

95

+

Figure 4.9 TADA1 complementation of hcef2. vH /LEF of intact leaves of Col-0 (filled circles)
and hcef2 P35S:!NTADA1 (open squares). Mean ± SD, n = 3.

96

85% (197). Both of these mutations result in partial loss of ribosomal proteins and impaired
+

chloroplast translation (197). The extents of CEF as measured by vH /LEF were increased by
about 2-fold compared to Col-0 in both prsp3 (0.0033 ± 0.0005 and 0.0017 ± 0.0001,
respectively, p < 0.001, n = 3) (Figure 4.10), and rps17 (0.0017 ± 0.0001 and 0.0055 ± 0.0004,
respectively, p < 0.001, n = 3). These results suggest that elevated CEF may be a general
response to disruption of chloroplast translation.
4.3.I Mass spectrometry proteomics
We used high-resolution proteomics (see Materials and Methods) to assess the effects of
hcef2 on the accumulation of both nuclear and plastid-encoded chloroplast proteins (Figure
4.11). Figure 4.11 shows statistically significant fold changes from Col-0 (log2, p < 0.05) of
thylakoid proteins in hcef2 grouped by complex. Each component of the electron transport chain
had significant changes for two or more subunits.
Strikingly, the relative changes of several thylakoid complexes were non-stoichiometric,
i.e. the changes in individual components of a complex differed from one another (Figure 4.11).
This effect is highly interesting because it should result in the accumulation of partially
assembled, and in many cases non-functional or dysfunctional, complexes. In photosystem II, the
PSBO2 (an isoform of the 33 kD protein of the oxygen evolving complex) (198) and PSBR
(involved in stabilizing the oxygen evolving complex) (199), proteins were diminished while
PSBF (the cytochrome b559 protein) was strongly increased (p < 0.01, n = 3).The PETB subunit
of the cytochrome b6f complex showed much higher accumulation than two other subunits of this
complex, PETA and PETC (p < 0.01, n = 3). Non-stoichiometric changes were also observed in
the accumulation of PS I antenna components, with non-stoichiometric decreases in PS I antenna

97

+

Figure 4.10 Elevated CEF in the ribosomal mutants prsp3 and rps17. vH /LEF of intact
leaves. Mean ± SD, n = 3.

98

Figure 4.11 Changes in the thylakoid proteome of hcef2. A) Fold changes from mean Col-0
values (log2) in the thylakoid proteome of hcef2. Mean ± SD, n = 3. B) Model of the thylakoid
membrane. Column colors of (A) correspond to the complex they are associated with in (B)10. *
p < 0.05, ** p < 0.01

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
"#!$%&!'()(!*+&'!),!-&.&/()&!01-*/&!23""!!4&/&!,5)(1.&'!56!718!73!91:+,.!(.'!;(/6!<1=),.!
()!)%&!Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory,
Richland, Washington 99354, USA
!

99

components LHCA1 and LHCA3 (p < 0.01, n = 3), while LHCA4 increases (p < 0.05, n = 3).
Likewise, the ATP synthase subunit ATPA decreased, while ATPE increased (p < 0.01, n = 3).
Finally, we also observed increases in relative abundance of all measured NDH subunits
(NDHA, NDHE, NDHO and NDH48, Figure 4.11), but the extent of accumulation varied
substantially (p < 0.05, n = 3).
4.3.J Flash induced relaxation kinetics in dark-adapted leaves of hcef2
The decay of the flash-induced ECS signal has been used to monitor the generation of
electric field across the thylakoid and its dissipation by the activity of the ATP synthase and
counterions (129). In dark-adapted leaves, or leaves infiltrated with methyl viologen (Figure
4.12), the ATP synthase becomes inactivated by oxidation of regulatory thiols, slowing the decay
of the ECS signal. The residual decay, measured at low flash intensity to prevent re-activation of
ATP synthase, reflects leakage of protons and counterions across the thylakoid membrane. In
Col-0, this residual decay was slow, with a lifetime of about 0.8 s, similar to previously
described results (129). A substantially increased ECS decay rate was about two-fold faster in
hcef2 (lifetime = ~0.4 s, Figure 4.12), indicating an increased rate of proton or ion leakage from
the lumen.
4.3.K hcef2 and related mutants show elevated H2O2 production
Figure 4.13 shows relative leaf H2O2 content in Col-0, hcef2, tada1, hcef2
P35S:!NTADA1, prsp3, and hcef1. Both hcef2 and tada1 had significantly higher H2O2
accumulation than Col-0 (2.67 ± 0.57, 1.67 ± 0.29, and 1.00 ± 0.06, respectively, p < 0.01 and p
< 0.02 respectively, n = 3) while the complimented line, hcef2 P35S:!NTADA1, had H2O2
levels similar to Col-0 (1.13 ± 0.16 p = 0.25, n = 3). The plastid ribosomal mutant prsp3 also had
100

Figure 4.12 Flash induced relaxation kinetics of the electrochromic shift in Col-0 (black)
and hcef2 (gray) leaves infiltrated with 100 µM methyl viologen. Data is representative of 3
independent experiments.

101

Figure 4.13 Relative accumulation of H2O2 in Col-0 and mutant lines. H2O2 levels were
assayed by resorufin fluorescence and normalized to chlorophyll content of the sample and then
normalized to average Col-0 fluorescence. Mean ± SD, n = 3. Asterisks indicate statistical
difference from Col-0.

102

a significantly higher level of H2O2 (1.41 ± 0.13, p < 0.01, n = 3). In addition, the hcef1 mutant
also was shown to accumulate increased levels of H2O2 than Col-0 (1.87 ± 0.30 p < 0.01, n = 3).
4.4 Discussion
4.4.A Disruption of protein translation in hcef2 leads to accumulation of H2O2 and
activation of CEF involving the chloroplast NDH complex
The hcef2 mutant was identified using a progressive screening approach as having
constitutively activated CEF. Several independent approaches demonstrated that hcef2 exhibits
strong activation of CEF, including comparison of proton and electron fluxes (Figure 4.4A), pmf
related parameters (Figure 4.4B), and postilumination fluorescence yield changes (Figure 4.5).
The elevated CEF in hcef2 was found to be insensitive to antimycin A, which inhibits the
PGR5/PGRL1 associated ferredoxin:plastoquinone reductase pathway (FQR) (87, 115) (Figure
4.6) arguing against the participation of the FQR pathway. Instead, the strong increase in
postillumination fluorescence rise and increased expression of the NDH complex (Figure 4.11)
suggests activation of the NDH pathway in hcef2.
4.4.B The hcef2 mutation disrupts chloroplast translation and leads to mis-assembly of
chloroplast protein complexes
The hcef2 gene was mapped to the TADA1 locus (At1G68720), coding for tRNA
Adenosine Deaminase Arginine, which is important for efficient chloroplast translation (195,
196). This assignment is supported both by observation of a similar phenotype in the knockout
mutant, tada1 (Figure 4.8), and by the loss of phenotype when hcef2 was complimented with the
C-terminus of the protein, containing the active site [Figure 4.9, see also (196)]. The C>T
transition in hcef2 introduced a nonsense mutation at R643, and effectively deleted the active site
of the enzyme (Figure 4.7), thus decreasing the ability of the chloroplast translation machinery to
103

recognize Arg codons CGC and CGA via the wobble mechanism of tRNA pairing (196).The
mutant must instead rely on the less efficient ‘two-out-of-three CGN’ codon recognition (196). It
was previously shown that disabling TADA1 affects the efficiency (or rate) of translation, but
does not impact its fidelity, i.e. proteins are made more slowly in tada1, but the amino acids are
inserted correctly (195, 196). This slowing of translation is likely the primary cause of decreased
growth in both hcef2 and tada1.
Strikingly, our proteomics results show that partially inhibiting plastid translation by
limiting availability of specific tRNA leads to alterations not only in the levels of photosynthetic
complexes, but in the stoichiometries of their component proteins. Of particular interest are nonstoichiometric changes in in subunits of PS II, the b6f complex, NDH and the chloroplast ATP
synthase (Figure 4.11). Because these complexes require a full complement of subunits for
function, hcef2 likely accumulates inactive or partially active complexes.
We also observed large changes in the stoichiometries of antenna complexes that appear
to alter photosynthetic responses. For example, an increase in the accumulation of PSBS, as seen
in hcef2 (Figure 4.11), is expected to increase the sensitivity of qE to pmf (24), as observed in our
results (Figure 4.3C). Likewise, a there was a alteration in the relative accumulation levels of PS
I antenna and core complex in hcef2 (Figure 4.11), which may explain the appearance of a 77K
fluorescence emission peak at 735 nm, and suggests loss of attachment of LHCI to PS I (200),
supporting our hypothesis that complex formation is impaired in this mutant.
By extrapolation, we propose that partially assembled redox enzymes, e.g. the
cytochrome b6f complex, would very likely result in production of reactive oxygen species. We
also observed altered ATP synthase subunit stoichiometric with ATPA decreasing and ATPE

104

increasing (Figure 4.11). Altered stoichiometry of this complex could result in an uncoupled
thylakoid membrane, as also suggested by the more rapid decay of the flash induced ECS signal
of dark-adapted leaves (Figure 4.12). Loss of protons through leakage or slip would disrupt the
output balance of ATP/NADPH resulting in inhibition of downstream metabolic reactions and
require the activation of additional ATP production processes, including CEF (2).
In chloroplasts of wildtype cells, disrupting one integral protein of a complex typically
leads to loss of all subunits, i.e. the expression and stability of the subunits are high regulated to
prevent the buildup of partially assembled complexes (100, 201, 202). In hcef2 this regulation
appears to be violated, leading to severe consequences for the function of photosynthesis. The
effect on subunit stoichiometry may be caused by the overall slowing of translation in hcef2 of
the CGC and CGA codons for arginine. The distribution of these codons within the chloroplast
genome is not even, and the efficiency of translation of a transcript is likely a function of CGC
and CGA codon content within the transcript. In effect, translation in hcef2 may not be slowed
evenly across the entire proteome, which may lead to a lag time during assembly and
accumulation of long-lived intermediates in complex assembly.
This hypothesis is supported by the hyper-accumulation of PETB protein in hcef2 (Figure
4.11). PETB attachment to subunit IV is the first step in assembly of the b6f complex (203)
followed by attachment of the Rieske protein (PETC). This may also explain stoichiometric
amounts of PETA and PETC in hcef2. Cytochrome b559 (PSBF) is thought to be part of an early
subcomplex in PS II assembly (204), and this protein accumulates at a higher level in hcef2,
while other PS II subunits decrease (Figure 4.11).

105

4.4.C Modifying chloroplast translation leads to accumulation of H2O2, a likely activator of
CEF
While the effect of the hcef2 mutation is certainly pleiotropic, likely affecting many
processes related directly and indirectly to photosynthesis, it could lend some insight into
regulation of CEF. For example, is was previously shown that simply suppressing overall
photosynthesis does not, by itself, trigger high rates of CEF (70, 78). Instead, CEF appears to be
dependent on the redox state of the chloroplast (69, 109, 119, 120), leading us to propose that
activation of CEF in hcef2, tada1 and related mutants may involve altered redox regulation.
Several lines of evidence suggest that this redox regulation may act through, or be
modulated by, the reactive oxygen species H2O2 (70, 99, 102, 119) (see Chapter 3). Several
mutants of the CBB cycle have found to have high CEF (69, 70, 79), all of which show elevated
H2O2 levels, and production of H2O2 within the chloroplast leads to a rapid increase of CEF in
vivo (Chapter 3).
The NDH complex has previously been shown to increase both accumulation and activity
in response to H2O2 (99, 102), suggesting a role for the observed increases in H2O2 in hcef1
(Figure 4.13) in the increased NDH activity in hcef2, as previously proposed (69, 104).
Because the codons affected in hcef2 (and tada1) are present in >80% of the protein
coding genes within the chloroplast genome (205), the overall effect on plastid translation is
expected to similar to that of prsp3, and rps17, but through a different mechanism. In other
words, the fact that hcef2, tada1, prsp3, and hcef1 all showed both increased levels of H2O2
(Figure 4.13) and elevated CEF (Figures 4.4, 4.8, and 4.10), suggests a model in which defects in
chloroplast translation lead to discoordination of protein homeostasis, leading to elevated H2O2
106

thus activating CEF. Finally, our results have immediate implications for any attempt to screen
for high CEF mutants as they may also identify lesions in the translational (and related)
machinery (as seen in the ribosomal mutants prsp3 and rps17, (Figure 4.10) in addition to
defects that directly involve photosynthetic processes.
4.5 Methods
4.5.A Plant materials and growth
All plants were grown photoautotrophically on soil in a controlled growth chamber with a
-2 -1

16:8 light/dark photoperiod (~100 µmols photons m s , white light). Seed for tada1 (GK119G08) and the tada1 line complimented with P35S:!NTADA1 was graciously provided by
Dr. Jo&e Gualberto. Seed for prsp3 (Salk_010806) and rps17 (Salk_066943) were provided by
the ABRC. The tada1 insertion was verified as described in (196). The hcef2 mutation in the
tada1 locus was verified by sequencing. The presence of the P35S: !NTADA1 construct was
verified using primers for the 35S promoter (5’-CCACTGACGTAAGGGATGACG-3’) and the
C-terminus end of TADA1 (5’-TGCTTTAGAACCCTCTCGAAT-3’). Verification of
homozygous prsp3 and rps17 was performed using primers generated from the SIGnAL T-DNA
primer design tool (http://signal.salk.edu/tdnaprimers.2.html).
4.5.B Isolation of hcef mutants
hcef2 was initially identified and isolated as a high NPQ mutant as described in (69).
Identification of backcrossed lines and F2 mapping populations with high NPQ was performed in
a high-throughput fluorescence-imaging chamber as described in (J. Cruz, L. Savage, R.
Zegarac, W. Kovac, C. Hall, J. Chen, R. Last, D. Kramer, submitted).

107

4.5.C 77K fluorescence spectroscopy
Fresh light adapted leaf material was flash frozen in liquid nitrogen, ground to a fine
powder and diluted to <5 µg/ml in ice as described in (206). Emission spectra were detected
using a spectrofluorimeter (Ocean Optics, HR200+ES) by a blue (440 nm) diode laser, controlled
by SpectraSuite software (Ocean Optics). The spectra were normalized to the 735 nm peak.
4.5.D In vivo spectroscopy
All in vivo spectroscopic measurements were performed on fully expanded leaves in
mature plants just prior to bolting. Comparisons were made between mature leaves, despite age,
due to the inhibited growth in the mutant lines. Steady-state chlorophyll a fluorescence yield and
light-induced absorption changes were made as extensively described elsewhere (7, 32, 35, 69,
70, 179, 187) on a spectrophotometer/fluorimeter described in (166). To account for changes in
pigmentation of the mutants, LEF was calculated using the approach of (69, 193) using the
following equation:

Eq. 4.1
!"# ! ! !!! ! ! ! ! ! !!!

where i is the actinic light intensity and A is the absorptivity of the leaf quantified using an
integrating sphere described in Idle and Proctor (207).
Electrochromic shift measurements were corrected for changes in leaf properties by
normalizing to leaf chlorophyll content. This correction gives similar results as corrections
described in (35, 69, 70).

108

For chlorophyll a fluorescence yield in response to short actinic pulses, plants were darkadapted for 10 minutes prior to the experiment. For each trace the excitation light was pulsed at a
frequency of 500 Hz, contributing minimally to the kinetics of chlorophyll a fluorescence
induction in the absence of actinic illumination. 5 actinic flashes (10 msec at ~12,000 $mol
-2

-1

photons m sec ) were given 0.2 s apart. A second experiment was performed in which an
interval of far red illumination was inserted after the last actinic flash. Data was normalized to F0
and FM, and the baseline was set to 0.
4.5.E Map-based cloning of hcef2
The hcef2 mutant was mapped on chromosome 1 between At1G68560 (25,733,701bp)
and At1G69020 (25,947,401), a 213.7 kb region, using molecular markers based on Simple
sequence length polymorphisms (SSLPs) and cleaved amplified polymorphic sequences (CAPS)
(208). Polymorphism sequence information between Col-0 and Landsberg erecta, from the
ABRC TAIR website (http://www.arabidopsis.org/browse/Cereon/index.jsp), was used to design
SSLPs and/or CAPS marker for mapping. F2 plants were derived from breeding homozygous
hcef2 (Col-0 background) and wildtype (Landsberg erecta background). The hcef2 mutation was
found to be recessive, and genomic DNA was isolated from homozygous F2 plants (hcef2 hcef2)
with high NPQ by chlorophyll a fluorescence imaging (described above). To determine the hcef2
mutation, we performed whole genome sequencing on homozygous hcef2 plants. Genomic DNA
from the mutant line was used to create a sequencing library using the Illumina TruSeq DNA
Library Kit following manufacturers recommendations. The library was then sequenced on an
Illumina GAIIx using single end with 50 base reads. All next generation sequencing was
conducted by the Genomics Core of the Research Technology Support Facility at Michigan State
109

University. Illumina reads were assembled using SeqMan NGen software (DNASTAR). SNPs
were compared across multiple samples of the same Col-0 background and SNPs unique to hcef2
were confirmed by Sanger sequencing.
4.5.F CAPS marker for hcef2 genotype
To genotype hcef2 without sequencing we designed a CAPS marker for the hcef2
mutation. A PCR fragment spanning the mutation site was amplified (5’GAGGCTGATTGGTCAAGGA-3’, forward; 5’-GGATGTTCAAAGGCTGTGGT-3’, reverse)
and then digested with NruI (R0192, New ENGLAND BioLabs). The Col-0 sequence is cut with
NruI, and homozygocity/heterozygocity was determined by comparing banding patterns on a 2%
agarose gel.
4.5.G Infiltrations
Freshly detached leaves were infiltrated with 20 µM antimycin A in distilled water in the
dark by soaking between 2 saturated lab tissues for 3 hours. Successful infiltration of antimycin
A was confirmed by secondary effects of this chemical on NPQ responses probed by chlorophyll
fluorescence (189). Leaves were infiltrated with 100 µM methyl viologen in a similar manner for
1 hour prior to measurement.
4.5.H H2O2 quantification
To quantify relative H2O2 accumulation, leaves were flash frozen in the light and H2O2
was extracted in 50 mM potassium phosphate (pH 7.4). Total H2O2 of the extract was quantified
using resorufin fluorescence, a byproduct of the reaction between Amplex Red (Invitrogen) and
H2O2, as described in Chapter 3.
110

4.5.I Determination of protein changes in hcef2
Total protein was extracted from whole leaf tissue as described in (69) and quantified
using a combination of SCX, LC, ESI, and MS/MS and analyzed using the peptide identification
software SEQUEST (209) in conjunction with the annotated protein translations from the
genome sequence of Arabidopsis thaliana TAIR 10 annotation (http://www.arabidopsis.org/).
Detailed methods are described in Appendix 1.
4.6 Author contributions
All data presented, except where noted, were obtained by Deserah D. Strand. Mapping of
the hcef2 mutation was started at WSU by Aaron K. Livingston, Heather M. Enlow, and Deepika
Minhas under the direction of Amit Dhingra. Mapping was finished by Mio Satoh-Cruz and
Tyson Koepke. Extensive preliminary data, and a preliminary version of the manuscript was
generated by Aaron K. Livingston (104). For the proteomics experiment, Kaori Kohzuma
designed the biological samples and replicates, and Kim K. Hixson performed the experiments
under the direction of Mary Lipton at PNNL. Kim K. Hixson and Mary Lipton analyzed the raw
data and transformation of the proteomics experiment, and assisted with the analysis and
interpretation of the final data sets.
4.7 Acknowledgements
The authors would like to thank Drs. Ralph Bock, Nicholas Fisher, and Jeff Landgraf for
helpful discussions. Experiments performed at MSU and WSU were funded by Grant DE-FG0211ER16220 from the Photosynthetic Systems program from Division of Chemical Sciences,
Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy
(to DMK). The Department of Energy Office of Biological and Environmental Research
111

Genome Sciences Program supported the high throughput proteomics component of this research
under the Pan-omics project. Work was performed in the Environmental Molecular Science
Laboratory, a U.S. Department of Energy national scientific user facility at Pacific Northwest
National Laboratory (PNNL) in Richland, WA. We would also like to thank our outstanding
technicians, Amelia Barhanovich, Brendan Johnson, Ryan Vink, and Ben Wolf for their
maintenance of plants and general assistance within the lab.

112

Chapter 5
Evidence for a proton pumping plastidic NAD(P)H dehydrogenase complex involved in cyclic
electron flow around photosystem I
Deserah D. Strand, Nicholas Fisher, and David M. Kramer

113

5.1 Abstract
Photosynthetic cyclic electron flow around photosystem I is thought to help balance the
ATP/NADPH energy budget of the chloroplast. In order to do this, it reroutes electrons from the
accepter side of photosystem I to reduce the plastoquinone pool. The proposed mechanisms of
plastoquinone reduction allow for, not only additional ATP production, but an alteration in the
+ -

H /e stoichiometry of the process itself. In this paper we show evidence that CEF in vivo
through the thylakoid NADPH dehydrogenase complex (NDH), homologous to respiratory
Complex I, has proton pumping function. This may allow for increased efficiency of ATP
+ -

production via CEF by increasing the H /e of CEF from the 2/1 expected of a non-proton
pumping route, to above 3/1, and may explain the low concentration of the NDH complex within
the thylakoid. In addition, a proton pumping NDH may allow a reversible process in which pmf
is consumed to oxidize plastoquinol and reduce NADPH, further adjusting the output of the light
reactions to meet downstream metabolic demands.
5.2 Introduction
The proton and electron circuits of photosynthesis are tightly coupled to each other. The
textbook ‘z-scheme’ model of linear electron flow, perhaps the best understood electron transfer
pathway in the thylakoid, provides a fixed number of protons transferred into the lumen,
generating proton motive force (pmf) for ATP synthesis, as electrons are passed through the
photosynthetic machinery. This ratio of electrons to protons is only altered if there is a slip or
leak in the reactions or alternative electron pathways are activated. Slip or leak is detrimental to
the organism because it has the potential to create reactive oxygen species (in the case of
electrons) or decrease the efficiency of photosynthesis (in the case of protons). Alternative
electron pathways in the chloroplast seem to be a more preferred option to the thylakoid, and
114

there are many that have evolved in plants, such as the plastid terminal oxidase, the water-water
cycle, the malate shunt, chlororespiration, and cyclic electron flow around photosystem I (CEF)
(41, 50, 59, 151) (discussed in Chapter 1). CEF as a process is interesting in that it seems to
involve multiple redundant pathways, and knocking out any one of them does not result in much
of a growth phenotype except under stress conditions (87, 100, 114). The best understood
pathway, at least structurally, is the thylakoid NADPH dehydrogenase complex (NDH), which is
analogous to respiratory Complex I. Another pathway is the antimycin A sensitive,
PGR5/PGRL1 associated pathway, in which it has been proposed these proteins form a complex
which is able to directly reduce plastoquinone to plastoquinol (PQ/PQH2) from electrons donated
from ferredoxin (Fd) (82, 83, 112, 113).
It is interesting that plants have conserved both of these routes of alternative electron
transfer, as they both are able to reduce PQ, and seem to have the same donor, Fd (82, 113, 167).
However, structurally we may have a clue to why these two are maintained. A complex of PGR5
and PGRL1 would likely only pass electrons to PQ, and any pmf generated from this process
would come from plastoquinol oxidation at the Qo site of the cytochrome b6f complex (b6f),
+ -

therefore the H /e stoichiometry of this route of electron transfer would be 2/1. On the other
+ -

hand, if the NDH is functionally similar to Complex I, it has the potential to increase the H /e

stoichiometry of CEF by pumping protons. The actual number of protons pumped per electron in
Complex I is still debated, but the consensus is 2/1 [reviewed in (210)]. If NDH maintains this
ratio, then NDH catalyzed PQ reduction, coupled with PQH2 oxidation by the b6f complex, the
+ -

H /e stoichiometry of this route of CEF could be as high as 4/1 (14, 15).

115

Based on several lines of evidence, including a high-resolution crystal structure, a
consensus model of Complex I proton pumping has emerged in which the reduction of the N2
iron-sulfur cluster leads to a conformational change of the membrane subunit NUOL, a
transmembrane proton channel which contains 2 C-terminal transmembrane alpha helices that
cross three additional transmembrane proton channels. This is thought to move polar residues
buried within each of the proton channels, releasing up to 4 protons (1 proton per channel) into
the intermembrane space (98, 210). It should be addressed that this proton number is still
+ -

debated, however the consensus from experimentation is that the H /e of Complex I is two
protons pumped for each electron passed to the quinone acceptor [reviewed in (210)].
In this work we show data supporting the hypothesis that NDH also acts as a proton
pump. We propose that the apparent redundancy in the routes of CEF exist in order to further
fine-tune the light reactions of photosynthesis to balance the chloroplast energy budget.
5.3 Results
A sequence alignment (Clustal Omega, Jalview) of Arabidopsis thaliana, Spinacia
oleracea, and Nicotiana tabacum single subunit membrane fraction NDH subunits with the
membrane fraction subunits of Complex I from Escherichia coli, Thermus thermophilus,
Yarrowia lipolytica, and Bos taurus, shows conservation of the indispensible polar residues E144
and K234 (NUOM numbering, Figure 5.1) in all analogous proton channels of NDH.
To test proton pumping capacity of NDH, we used the hcef1 mutant, which shows an
increase in relative CEF through the NDH complex (69). Using changes in absorbance of 820 nm
during a dark interval, we can calculate relative electron transfer rates through PS I (vP700).
+

Comparisons of transthylakoid proton flux (vH ) and electron transport rates through PS I and PS

116

026789:;<=>@TB!"#!$!%!"&'%'(&)!"!%*"!%)('!+,'!"""'-./0$1111')-!0(/!*%!%*'!(/#!,
0267*CDD9E;C>??UB+!("/!+"!!0%.(!2*"#%*"!%)('!,!!%"!"3"*/##111121#)-(#%*+!&%'!#/#!%
0H9?I7'J=DKLM>@TB(!!&)/!!!/!%((-2!!+%*+%%)((!",(!!&!*!*//11111)/-'-(!*'%+!%'!+/#!,
0L9&7)N9O;>??@B!&."!//+"/+%!("2&%!%%%%.)&&!+,&*%!"(!./34(#2/4'-"'(('4%%"*'$(#/!+
0PJ27CDCQJ>??UBR&!(&*#/$"/#%##-2!!!%%"&.)!)!",+*!!!#&.//1111144-!*#('4%"!*'(/##"%
0PJ27MS;<CNJ>??UB &!(&*#($"/!%##-2!!!%%"&.)!)!",+*!!!#&.//1111144-!*#('4%"!*'(//#"%
0PJ27Q98CN>??UB &!(&*#/$"/#%##-2!!!%%"&.)!)!",+*!!!5&.//1111144-!*#('4%"!*'(//#+%
'&6

'&A

'&V

026789:;<=>?@AB .!(5&&(%&+4&,!*/!3!.!,4(3+)(,"(/#&+!((+!!4!
0267*CDD9E;C>F?GB .!/!%"("&+4',!%,"3+.!,++3#)#,!(/#&"!(/!"!4!
0H9?I7'J=DKLM>?@IB%!/%(!(%("4',!%,!3(.!,,%3$)03,#/!(2(!/'!*4+
0L9&7)N9O;>FFIB&!/%%"(%(+4&,++,!3/.!,2(3#$(,'(/#+2!(/"!!4'
*"/%!"(%(+4#,"",!3'.!,2'3/)(3*#'5&!!(/"!!4&
0PJ27CDCQJ>F?IBR
0PJ27MS;<CNJ>F?IB*"/%%"(%(+4!,"",!3'.!,2'3/)(3*#'5&!!(/"!!4&
*"/%%"(%(+4#,"",!3'.!,2'3/)(3*#'5&!!(/"!!4&
0PJ27Q98CN>F?IBR
'&U

'&T

Figure 5.1 Residues essential for proton pumping are conserved in the NDH membrane
fraction. Representative proton channel alignment of plant NDH subunits with Complex I.
NdhD is analogous to ND4, Nqo13, and NuoM in respiratory Complex I11. Sequences are from
the UniProt database for Bos taurus (bovine), Yarrowia lipolytica (Yarrowia), Thermus
thermophilus (Thermus), Escherichia coli (Ecoli), Arabidopsis thaliana (arath), Spinicia
oleracea (spinach), and Nicotiana tabacum (tobac).

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
11
Figure 5.1 was kindly generated by Dr. Nicholas Fisher.
117

+ -

II, we can calculate the H /e stoichiometry for CEF in this mutant. In Figure 5.2A we see an
increase in vP700 of ~50% in hcef1 when compared to Col-0, (m = 0.19 +/- 0.04 and 0.135 +/0.011, respectively, p = 0.0002, n = 3), while in Figure 5.2B we see a nearly 4-fold increase in
+

vH as a function of LEF in hcef1 when compared to Col-0 (m = 1.28 +/- 0.24 and 0.26 +/- 0.037,
+ -

respectively, p < 0.0001, n = 3). This suggests H /e stoichiometry in excess of the 3/1 expected
+

of LEF. Additionally, Figure 5.2C shows an increase in the amount of vH to electron transfer
+ -

through photosystem I, indicating proton pumping CEF in hcef1. If the H /e of CEF in hcef1
were 2/1 as expected for a non-proton pumping route, we would expect a decrease in the slope of
this relationship.
To qualitatively test for proton pumping, we monitored ATP synthesis, using luciferase
phosphorescence, in the dark after addition of electron donors to the cuvette. There was a slow,
but reproducible, increase in slope, indicating ATP synthesis (Figure 5.2D), and therefore pmf
generation in the dark, which, since there is no electron sink to drive the b6f complex, could only
be explained by a proton pumping plastoquinone reductase. In addition, ATP synthesis was
insensitive to tridecyl stigmatellin, a potent inhibitor of the b6f complex, and oligomycin, a
mitochondrial specific ATP synthase inhibitor (data not shown), eliminating pmf contribution
due plastoquinol oxidation at the b6f complex or by mitochondrial contamination.

118

Figure 5.2 Evidence for involvement of a proton pump in CEF. A) Electron flux through PS I
(vP700) as a function of electron flux through PS II (linear electron flow, LEF). B) Transthylakoid
+

+

proton flux, vH , as a function of LEF. C) vH as a function of vP700. D) ATP synthesis,
monitored by luciferase luminescence, in DCMU treated spinach thylakoids in the dark after
addition of decylplastoquinone (dPQ)12.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
12
Figure 5.2D was graciously supplied by Dr. Nicholas Fisher.
119

5.4 Discussion
5.4.A CEF involves a proton pump
Using the Arabidopsis mutant hcef1, which is proposed to have high rates of CEF
primarily through the NDH complex, we have shown the first evidence of a proton pumping CEF
in vivo. The hcef1 mutant accumulates high levels of NDH dependent CEF. There have been
multiple mechanisms of CEF proposed in higher plants and algae. Of these known pathways, the
only protein complex that may be structurally capable of proton pumping would be the NDH
complex. Despite its usually low expression levels, the proton pumping activity would allow for
higher ATP generation via CEF than the other proposed pathways.
The NDH complex is homologous to Complex I, and evolution has conserved many of
the residues proposed to be involved in the mechanism of proton pumping. The apparent H+/estoichiometry for CEF in these conditions is above 3/1, which is in agreement with a >1/1
stoichiometry of a Complex I like NDH assuming a b6f ratio of 2H /1e is maintained (14).
+

%

It should be mentioned that the proton pumping capability of NDH must satisfy
+ -

thermodynamic constraints. At equilibrium, the relationship between protons pumped (H /e )
into the lumen against the pmf (!p) per electron, with a redox span of !Eh mV is given by:

Eq. 5.1
!!! ! ! !!!

+ -

where n represents the H /e ratio. With Fd as electron donor to NDH, !Eh equals 518 mV
assuming 90% reduction of the PQ and Fd pools respectively. Assuming a !p of -180 mV across
120

+

-

the thylakoid membrane in the light, NDH would be capable of acting as a 4H /2e pump. This
assertion also holds for a predominantly oxidised (90%) Fd pool. If NADPH is the electron
donor to NDH the energetics are more constrained, with a !Eh of 380 mV under conditions of
+

-

90% reduction of the NADPH and PQ pools. 4H /2e pumping is possible only up to a !p of 190 mV in this instance. In vivo it has been estimated that under light saturating conditions
+

-

maximum pmf is ~175 mV (26), a value that still allows for the forward reaction of a 4H /2e

proton pumping NDH complex. Mitochondrial Complex I is confronted by this problem during
respiratory state 4 (ADP exhausted) when the pmf force may approach -220 mV, and there is
+

-

insufficient redox energy for the enzyme to operate as a 4H /2e pump (211). Under these types
of conditions we would not expect a need for NDH to operates as this would likely represent an
ATP surplus, and may lead to the reverse reaction of the enzyme (discussed below).
This observation may explain why there are multiple routes of differentially regulated
cyclic electron flow in chloroplasts (reviewed in Chapters 1-5). We propose a model in which the
two most studied routes of CEF, in higher plants, are activated step-wise when the downstream
metabolism decreases ATP/NADPH. First, a reducing environment is generated as NADPH
accumulates, and CEF activated through the presumably non-proton pumping, rapidly reversible
antimycin A sensitive pathway (see Chapter 2, Figure 2.5A). Under conditions in which this
route of CEF is not able to augment the ATP deficit, and restore redox homeostasis to the
chloroplast, ROS is generated and activates a proton pumping NDH complex [see chapter 3,
-

Figure 3.9 and (99, 102)]. This would increase ATP formation per e transfer, increasing the
efficiency of ATP production via CEF. This route of CEF is not as rapidly activated as the FQR

121

(see chapters 2 and 3), and long term ROS generation leads to not only activation of already
assembled complexes, but an increase in total NDH content (99, 102), likely indicating slower
reversibility of this response.
5.4.B A proton pumping NDH would be reversible
The discovery of a proton pumping NDH has further implications for energy balance
within the chloroplast in that a proton pumping NDH should also be reversible, consuming pmf
+

to oxidize PQH2 and reduce NADP to NADPH. Using several assumptions in relation to the
+

-

+

mechanism of a reversible NDH, including a reaction of 4H /2e and NADP as an acceptor, we
can calculate the free energy of the reverse reaction (!Grev) of NDH under a variety of
conditions. Figure 5.3 shows !Grev (25°C, pH 7.5) as a function of pmf at different redox states
+

of the donor (PQH2) and acceptor (assumed to be NADP ). The reaction is favored with
increasing pmf and with increasing reduction of the PQ pool and oxidation of the NADPH pool.
We might expect reversibility of this process to be physiologically important during
conditions where NADPH is limiting, but pmf and plastoquinol are high. This situation may
occur during induction of photosynthesis where reducing equivalents (Fd, NADPH) are being
consumed by the thioredoxin-dependent activation of sink metabolism and ATP synthase,
leading to a NADPH deficit. Inactive ATP synthase could potentially lead to a high pmf and
slowed quinol oxidation at Qo, leading to an increase in PQH2/PQ. These conditions would favor
the reverse reaction of NDH, which may have a dual role in chloroplast energy balance as it
would consume pmf, and the potential for ATP production, to produce NADPH, altering the
ATP/NADPH of the light reactions (Figure 5.4).

122

Figure 5.3 Directionality of NDH. Conditions in which the forward reaction of NDH is
favorable (above dashed line), or the reverse reaction of NDH is thermodynamically favorable
+ -

(below dashed line)13. Calculations were performed assuming a 2H /e NDH.

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
13
Figure 5.3 was graciously supplied by Dr. Nicholas Fisher.
123

Figure 5.4 Hypothetical model of the reversible reactions of NDH. When !GATP is large, pmf
is correspondingly large, in such an instance, protons are consumed from the lumen to drive
+

oxidation of PQH2 and ultimately, reduction of NADP to NADPH.

124

It should be clarified that these calculations are made with many broad assumptions, many of
which, if violated, may lead to the complex being irreversible. First, it has been proposed that Fd
is the electron donor to NDH in the forward reaction (167), but if Fd is the acceptor in the
reverse reaction, pmf required to drive the reaction is calculated to be too high to be
physiologically relevant, and under these conditions the complex would be essentially
irreversible. Secondly, the pH of the quinol binding pocket of NDH is an unknown, and in Figure
5.3, assumed to be 7.5. Within the calculation of !Grev, as pH decreases, the amount of pmf
required to drive the reverse reaction increases, and could conceivably reach a point where the
reaction is irreversible.
5.5 Methods
5.5.A Plant materials and growth conditions
Arabidopsis thaliana plants were grown in a 12:12 light:dark photoperiod. Col-0 was
measured at 3-4 weeks of age. Slow-growing hcef1 was measured at the same developmental
stage as Col-0, which was around 6-7 weeks of age.
5.5.B In vivo spectroscopy
Variable chlorophyll a fluorescence was used to calculate the quantum yield of PS II ($II)
as described in (7, 187). In order to determine LEF rates, we took a more complex approach
expanded on in Appendix 2.
Redox state of PS I was monitored using absorbance changes at 820 nm in a protocol
modified from (68). Plants were poised by illuminating with 700 nm actinic light, and the initial
+

rate of P700 re-reduction kinetics were used as the relative rate of electron transfer through PS I
-2 -1

(vP700, !A820 nm m s ).

125

+

-2 -1

Transthylakoid proton flux (vH , !A520 nm m s ) was calculated using the
electrochromic shift of the carotenoids at 520 nm as described in (69) and Chapters 3 and 4. To
correct for variability in leaf pigmentation between Col-0 and hcef1 total extent of ECS was
normalized to the amplitude of the PS I contribution to ECS in the presence of DCMU (see
Appendix 2, Figure A2.4). This gave similar results to other methods of normalization described
in (69) and Chapters 3 and 4.
5.5.C Quantification of in vitro ATP production
Proton pumping in vitro was measured in the dark as ATP production using the Promega
(ENLITEN) luciferase/luciferin reagent kit with a laboratory-constructed PMT phosphoroscope.
Osmotically ruptured spinach chloroplasts (prepared in the presence of 10 mM DTT) were
present at a chlorophyll concentration of 40 µg/ml. DCMU was present at 10 mM. Fd and
NADH were present at 5 µM and 1 mM respectively. The assay buffer (pH 7.6) consisted of 10
mM HEPES, 10 mM potassium phosphate, 5 mM MgCl2, 10 mM potassium phosphate with 2
mM DTT and was supplemented with 10 mM ADP and 100 µM diadenosinepentaphosphate (an
adenosine kinase inhibitor). Proton pumping was initiated by the addition of 50 µM
decylplastoquinone (dPQ), and the proton gradient was collapsed by the addition of 10 µM
gramicidin. The plastid ATP synthase was activated by actinic illumination (625 nm) for two
minutes immediately prior to the start of data collection.
5.6 Author contributions
All data presented, except where noted, were obtained by Deserah D. Strand. Dr.
Nicholas Fisher optimized and performed the in vitro assay for ATP synthesis in the dark.

126

5.7 Acknowledgements
I would like to thank Dr. Nicolas Fisher and Dr. Jeff Cruz for helpful discussions. This
work was supported by Grant DE-FG02-11ER16220 from the Photosynthetic Systems program
from Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy
Sciences of the US Department of Energy (to David M. Kramer).

127

Chapter 6
Conclusions and future directions
Deserah D. Strand

128

6.1 Abstract
This chapter summarizes current consensus in the field in light of the data presented in
this dissertation, and discusses questions that remain open, or have emerged from the findings in
the previous chapters.
6.2 Conclusions
Throughout human history crop yields have been improved through trial and error with
traditional breeding and selection. As the world increases its demand for food and renewable fuel
sources, the need to understand plant and algal physiology and biochemistry increases in parallel.
We have entered an age of technology and human understanding of complex biochemical
processes, yet we have made very little progress in improving the photosynthetic reactions of
plants and algae, despite these reactions being some of the best characterized in all of biology.
Clearly we still have unanswered questions.
Many of these unanswered questions lie in the alternative electron pathways within the
chloroplast, which are present in much more abundance than in the mitochondria alternative
electron transport chain. As discussed in Chapter 1, some of these do not seem to contribute
significantly towards energy balance, but instead are important during development (plastid
terminal oxidase) (212), under prolonged environmental stress (malate dehydrogenase, NDA2)
(41, 43, 44, 117), or for redox poising of the PQ pool (NDA2) (86, 116). In light of the work
presently being carried out in the Kramer lab on mutants in chloroplast targeted genes with
unknown functions, it is likely the full extent of the roles of these alternative electron transport
proteins is yet to be discovered (J. Cruz, L. Savage, R. Zegarac, W. Kovac, C. Hall, J. Chen, R.
Last, D. Kramer, submitted). In line with this, CEF pathways could also be considered among
some of the last uncharacterized complex(es) in photosynthesis.

129

In light of recent work it may seem as if the FQR has been identified as a PGRL1-PGR5
complex that acts as a quinone reductase. However, this is not yet the case. As discussed in
chapter 1, there are major flaws in the enzymatic assays that are used to support this conclusion,
and therefore the identity of the antimycin A sensitive FQR is still yet undetermined. It is clear
that PGRL1 and PGR5 have a role in the antimycin A sensitive pathway of CEF in plants, but
they may be part of an even larger complex as is seen in algae, or may be regulatory as proposed
by Aro and coworkers (114). Chapter 2 discusses the rapid modulation of the FQR by redox
state, and suggests the involvement of a regulatory thiol. The Em of FQR activity is in agreement
with the Em of a thiol present on PGRL1, and supports a regulatory role of this complex as part
of, or in association with the FQR. A broader regulatory role for PGR5 is suggested by the large
defect in ATP synthase modulation in its absence, a defect that is more likely to explain the
dramatic loss of NPQ in this mutant than the loss of CEF. These distinctions are important,
because increased emphasis on the role of CEF could lead to an inaccurate model of the light
reactions and their regulation.
Despite extensive work on the characterization of NDH (84, 94–96, 167, 183, 213), there
are still many unknowns about this complex. There is difficulty in studying NDH, and this may
lie in the scarcity of the complex in unstressed Arabidopsis leaves (69, 95). With the recent
crystal structure of Complex I a complete structural set of the respiratory electron transport chain
has been obtained, and increasing mechanistic studies are possible (97, 98, 210, 214). In the
future we can expect similar discoveries with NDH, unlocking one of the last unknowns of the
photosynthetic electron transport chain. In the absence of a structure for NDH, we still have
discovered many things about this complex. Several groups have done extensive work on
identifying the subunits of NDH, and we now have identified >22 nuclear and plastid encoded
130

genes, as well as several that are plant specific (94). Ongoing research by a number of groups
continues to lend insight into assembly, association, and expression of NDH (95, 183, 215).
Functional understanding of NDH is complicated by the difficulty of assessing the
activity of the complex. Activity has been measured indirectly by ferricyanide reduction, in gel
reduction of NBT in the presence of NADH, and chlorophyll a fluorescence changes (84, 85, 99,
213). Quinone reduction assays have also been attempted with soluble quinones, but are likely
not physiologically relevant (85). These methods together have yielded new insights but have
also led to some key discrepancies (i.e. rates of electron flow) (84, 87, 163) and contradictions
(i.e. identity of electron donor) (85, 99, 167). It is clear there is a lot of work to still be done in
order to understand even the basic function of this complex.
While in vitro work seems to be the standard for NDH activity, the Kramer lab has an
increasing body of research on CEF activity in vivo. In this dissertation I have presented some
key data that begins to fill in the gaps of our understanding of CEF regulation. In addition to the
in vitro observations of FQR activity in response to redox state described in Chapter 2, Chapter 3
is the first description of CEF (via NDH) activation in response to ROS in vivo. These findings
are complimentary in that we see differential activation of the two independent pathways of
CEF, allowing a response to an ATP deficiency that could have the consequences of both a more
reduced stroma, or the buildup of reactive intermediates and subsequently ROS. We additionally
see differences in the rate of activation of the two patways, with the FQR rapid (seconds to
minute timescale, Chapter 2, Figure 2.4) and the NDH slower (minutes timescale, Chapter 3,
Figure 3.6). Additionally, Chapter 5 supports the NDH as a proton pumping quinone reductase,
indicating that the two pathways operate at different ATP generating efficiencies. This supports
the model proposed in Chapter 2 that the FQR is a ‘first response’ to ATP/NADPH imbalance,

131

and allows a more inclusive model of both CEF pathways as a way to fine tune alternative
electron transport in the chloroplast.
Chapter 5 also postulates the function of a reversible proton pumping NDH in the
thylakoid (Figures 5.3 and 5.4) in which an imbalance in the ATP/NADPH production with and
excess of ATP could be corrected by NDH consuming pmf to drive oxidation of PQH2 and
+

+ -

reduction of NADP . This would allow for the increased flexibility of bulk H /e transfer by the
electron transport chain.
6.3 Future directions
Obviously there are still many unanswered questions involving chloroplast energy budget
and specifically CEF. The models proposed within this dissertation are far from complete, and
require additional rigorous testing. Despite a clear role for ROS signaling in CEF (99, 102)
(Chapter 3), we still do not know much about the processes that lead from the signal to
activation. The inducible system introduced in Chapter 3 and (176) may serve as a valuable tool
in the elucidation of this pathway, and the next step from this work could be a suppressor screen
of the GO5 mutant. Previous work points to the potential role of Ca

2+

signaling and

phosphorylation in activation of CEF (102), and it may be that we find proteins involved in
either of these that when absent lead to the insensitivity of NDH to peroxide. With the
observations presented in (102) and Chapter 3, the field is very open for experimentation.
Likewise, determining the proteins involved in FQR-mediated CEF is the next step in the
study of this pathway. Evidence for PGRL1 acting as a direct quinone reductase is ambiguous
(discussed above), however, PGRL1 is a good target to look for a complex formation or
interaction with a protein that would serve as an appropriate quinone reductase. This has been
proposed to be the Qi site of the cytochrome b6f complex (b6f), as the b6f complex possesses an
132

additional heme Ci not seen in other bc complexes, and closely associates with FNR (110, 168).
It is possible that PGRL1 and PGR5 associate further with the b6f complex to facilitate or
activate electron transfer through the heme Ci, and similar associations have been proposed
previously (92, 109, 113). This is a difficult hypothesis to test, as we currently lack the ability of
inhibit the Qi site of the b6f complex, and quinone reduction would likely still occur with the Qo
site inhibited. The reverse genetics approach, despite the difficulties presented in chapter 4, has
proven to be quite useful in CEF research, as it has allowed the identification of several mutants
with relatively high rates of CEF. While a good proportion of mutants we have identified as hcef
accumulated high levels of H2O2, and as such are likely activated in the NDH pathway of CEF,
we have found a few that do not, and these may have increased rates through the FQR. If this is
the case, these mutants may provide valuable tools in which to study the antimycin A sensitive
pathway. While results are still very preliminary, I am very optimistic and excited to see what
comes of these studies.
Finally, as discussed in Chapter 5, the confirmation of a proton pumping NDH is a major
discovery in the field of chloroplast bioenergetics. This finding not only substantially increases
our structural and evolutionary understanding of NDH, it also increases our regulatory
understanding of photosynthesis. It allows for an integrated model of the multiple proposed
routes of CEF, and gives insight for why these seemingly redundant pathways have been
conserved in plants. The calculations presented in Chapter 5 also show a large gap in our
knowledge of NDH (discussed in Chapter 5). We know nothing about the quinone binding site of
NDH, we have contradictory evidence as to the identity of the donor (84, 99, 167), and we don’t
+ -

yet know the H /e of the proton pumping process. These unknowns lead to an incomplete model
133

as to contribution of pmf by the NDH and the conditions in which we would expect NDH to be
reversible, and it is very likely this model is not sufficiently descriptive. Therefore, the next steps
in this line of research are to characterize the NDH structurally and functionally. This is much
easier said than done, but with time and effort we may make the same extraordinary progress as
has been seen recently with respiratory Complex I.
Together, the findings within this dissertation are another step towards our complete
understanding of CEF and chloroplast energy balance. I look forward to what the scientific
community will learn in the next decade and beyond.

134

APPENDICES

135

Appendix I
Supplemental methods for chapter 414

!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
"2!These methods were supplied by Kim K. Hixson and Mary Lipton at Pacific Northwest
National lab, and will appear in the supplemental material of the published form of chapter 4.!
136

A1.1 Analysis of protein abundance
The following chemicals used, unless otherwise noted, were obtained from the SigmaAldrich Company (St. Louis, MO) and were of analytical grade. After extraction, protein was
separated into the soluble and membrane fractions and each sample was dried down using
evaporative centrifugation and was suspended in 200 mL of 7M urea, 2M thiourea, 4% CHAPS,
2 mL of TCEP in 50 mM ammonium bicarbonate, pH 7.8. Vortexing and sonication were used to
aid in solubilizing the pellet. Total protein concentration was determined by Bradford (Pierce,
Rockford, IL).
A1.2 Trypsin digestion and cysteine alkylation
Proteins were denatured and digested with trypsin as described elsewhere (216). 20mM
iodoacetamide was added and samples were then incubated at room temperature in the dark.
Samples were dried using evaporative centrifugation and resuspended in 1 mL of 25%
acetonitrile and 10 mM ammonium formate, pH 3.0 with vortexing brief sonication. Samples
were centrifuged at 13,500 x g for 5 min and the supernatant was used in subsequent steps.
A1.3 Peptide concentration and cleanup
The digests were desalted using Supelco (St. Louis, MO) Discovery 1mL 50 mg SCX
SPE tubes. Columns were washed with 1 mL of methanol followed by 2 mL of 25% acetonitrile
in 10 mM ammonium formate, pH 3.0. Samples were then loaded and washed with 6 mL of 25%
acetonitrile in 10 mM ammonium formate, pH 3.0. Samples were eluted with 5% NH4OH, 15%
water, and 80% methanol, dried via evaporative centrifugation, and resuspended in 60 mL of
nanopure water with vortexing and sonication. Samples were then centrifuged at 13,500 x g for 5
min the supernatant was taken. Total protein concentration was determined by Bradford, and
diluted to a concentration of 1 mg/mL. A peptide sequence library, containing peptide sequences,

137

elution time, and parent peptide mass, was constructed by MS/MS data acquisition using peptide
pools with equal mass from each sample.
A1.4 Strong cation exchange (SCX) fractionation of peptides for potential mass and time
(PMT) tag acquisition and capillary LC separations
300 mg of each peptide pool were separated with a SCX fractionation as described
elsewhere (217). All SCX fractions and 2.1 mg of each sample were individually separated by
an automated in-house designed HPLC system as described next. The HPLC system consisted
of a custom configuration of 100 mL Isco Model 100DM syringe pumps (Isco, Inc., Lincoln,
NE), 2-position Valco valves (Valco Instruments Co., Houston, TX), and a PAL autosampler
(Leap Technologies, Carrboro, NC), allowing for fully automated sample analysis across four
separate HPLC columns (218). Reversed-phase capillary HPLC columns were manufactured inhouse by slurry packing 3 mm Jupiter C18 stationary phase (Phenomenex, Torrence, CA) into a
60 cm length of 360 µm o.d. x 75 µm i.d. fused silica capillary tubing (Polymicro Technologies
Inc., Phoenix, AZ) that incorporated a 0.5 mm retaining screen in a 1/16” custom laser-bored 75
mm i.d. union (screen and union - Valco Instruments Co., Houston, TX; laser bore - Lenox Laser,
Glen Arm, MD). Mobile phase consisted of 0.2% acetic acid and 0.05% TFA in water (A) and
0.1% TFA in 90% acetonitrile/10% water (B). The mobile phase was degassed by using an inline Degassex Model DG4400 vacuum degasser (Phenomenex, Torrence, CA). The HPLC
system was equilibrated at 10 kpsi with 100% mobile phase A, and then a mobile phase selection
valve was switched 50 min after injection, which created a near-exponential gradient as mobile
phase B displaced A in a 2.5 mL active mixer. A 30-cm length of 360 mm o.d. x 15 mm i.d.
fused silica tubing was used to split ~20 mL/min of flow before it reached the injection valve (5
mL sample loop). The split flow controlled the gradient speed under conditions of constant
138

pressure operation (10 kpsi). Flow through the capillary HPLC column when equilibrated to
100% mobile phase A was ~500 nL/min.
A1.5 Peptide mass and time tag (PMT) acquisition
The elute from the HPLC was directly electrosprayed into an ion trap MS (LTQ,
ThermoFinnigan, San Jose, CA) using electrospray ionization (ESI). The mass spectrometer
operated in a data-dependent MS/MS mode and the peptides were analyzed with one full m/z
range (400-2000) each. The details for PMT generation are described elsewhere (219). A total of
152 peptide fraction and complex peptide sample runs produced MS/MS spectra which were
analyzed using the peptide identification software SEQUEST (209) in conjunction with the
annotated protein translations from the genome sequence of Arabidopsis thaliana TAIR 10
annotation (http://www.arabidopsis.org/). A dynamic modification search (i.e., the presence and
absence of the modification was searched) for methionine and proline oxidation and a static
search (i.e., presence of the modification was searched only) for alkylation of the cysteines by
iodoacetamide. Non-enzyme cleavage constraints were applied. Qualitative/cursory
identifications in the putative mass and elution time (PMT tag) peptide library were based on a
minimum cross correlation (Xcorr) score of 2 for all peptides identified at least twice in all
MS/MS experiments.
A1.6 Accurate mass and time (AMT) tag identification and alignment
Using 2.1 µg of total peptide from each individual time point sample, intact peptide mass
(MS) data were obtained using the same HPLC system but using a ThermoScientific Exactive
Orbitrap mass spectrometer (Thermo Scientific, San Jose, CA) outfitted with a custom built
electrospray ionization (ESI) interface. The electrospray emitter was custom made using 150
mm o.d. x 20 mm i.d. chemically etched fused silica column (220). The heated capillary

139

temperature and spray voltage were 200°C and 2.2 kV, respectively. Data were acquired for 100
min., beginning 65 min. after sample injection (15 min. into gradient). The mass spectrometer
was set to record spectra from m/z 400 to 2000 at a resolution of 100,000 and AGC setting of
6

3x10 . Each sample was run in duplicate. The run order for each run set of duplicates was
randomized in a Latin Squares design. A mass calibration mixture was infused at the end of each
analysis, and the masses of the compounds in the mixture were used to calibrate all of the spectra
within the analysis.
The MS data obtained from the Exactive MS were subsequently processed using the
PRISM Data Analysis System, a series of software tools developed in-house. The data were
initially de-isotoped to give a monoisotopic mass, charge, and intensity of the major peaks in
each spectrum. The data were then analyzed in a two-dimensional fashion to determine the
groups of peaks that were observed in sequential spectra.
Each group identified as a unique mass class (UMC) feature, was characterized as having
a distinctive median mass, central normalized elution time (NET) (221), and its abundance was
estimated by the maximum of the intensity for its MS peaks. Each UMC was determined by
comparing the mass and NET to those in the PMT tag database that passed the p-value cutoff of
0.01 using the in-house developed software Viper (http://omics.pnl.gov/software) (222). Filter
tolerances were set to ± 2 ppm for the mass and ± 2.5% for the elution time. Those UMCs (MS
only data from the Exactive) that most closely matched the PMT tags (MS/MS peptide data from
the LTQ iontraps) and whose predicted parent mass was verified by the high mass measurement
accuracy provided by the Exactive Orbitrap MS were validated as Accurate Mass and Time
(AMT) tags resulting in a list of peptides observed and an abundance value for each which was

140

calculated by Viper as the sum of the peak intensity of each eluting peptide over the time it was
observed for the most abundant charge state.
A1.7 Normalization of replicate analyses
Peptide data that resulted after peak matching and STAC_UP score filtering (> 0.5), were
then transformed into log2 values and were normalized using the mean center normalization
function in the publically available software Inferno
(https://code.google.com/p/inferno4proteomics/). Proteins with peptides shared among many
proteins were grouped together. Next all the peptides unique to one protein and the peptides that
are shared among multiple proteins, were “rolled up’ into a single protein or protein group value
by using the Rrollup function in Inferno. Rrollup works by first identifying the most abundant
and prevalent (seen in most LC-MS runs) peptide for each protein or protein group. Using the
abundance value profile of this peptide all other abundance profiles for all other peptides
belonging to that protein are scaled (normalized) to it. The protein value is then calculated by
taking the median value of all normalized peptides in that run. For proteins seen 5% of the time
in all datasets with 1 or more unique peptides identified, an additional Grubbs’ test (p-value
cutoff 0.1) was performed to eliminate outlier profiles from the rollup calculation. Proteins with
only 1 peptide identified were included for consideration and so the number of peptides
identified for each protein is included in all data tables so that those proteins can still be
considered but given less weight.
A1.8 Log fold change determination
Log2 values of the Rrollup protein/protein groups were transformed back to a non-log
transformed value by calculating 2^(log2 changes in protein abundance). Missing values were
imputed with 160 which was a value less than the minimum abundance value in the entire

141

dataset. Protein abundances determined to be significantly represented within the data set were
isolated for each protein complex of interest and positive and negative fold changes were then
determined for each hcef2 sample versus the average Col-0 abundance for either the insoluble or
soluble samples.

142

Appendix 2
Towards a more precise measure of electron transfer rates through photosystem II
Deserah D. Strand and David M. Kramer

143

A2.1 Abstract
The use of chlorophyll a fluorescence in photosynthetic measurements is considered the
gold standard in labs all over the world. However, the calculations for photosynthetic electron
transport often make many assumptions about the system being measured. This manuscript
discusses these assumptions and proposes conditions in which these assumptions are likely
violated. Specifically, the method for calculating PS II electron transfer rates, or linear electron
flow, is discussed, and a method for the careful correction of these rates is presented.
A2.2 Introduction
Chlorophyll a fluorescence yield is frequently used to calculate photosynthetic
parameters in a relatively simple and non-invasive manner. It has been used extensively in basic
research to identify and characterize structural and regulatory components of the photosynthetic
electron transport chain (7, 82, 84, 223). In addition to this, chlorophyll fluorescence imaging has
been used in a wide range of crop specific research, such as breeding efforts to identifying plants
with increased stress or disease resistance and in the mapping of beneficial QTLs (224–226)
[reviewed in (227)]. The current trend of research towards high-throughput screening methods
using this technique, in conjunction with fluctuating light or stress conditions, has allowed the
identification of conditions in which mutants in chloroplast targeted proteins show a fluorescence
phenotype when none was previously described, and has the potential to make a large impact on
our understanding of photosynthesis and how it relates to downstream metabolism and stress
response (J. Cruz, L. Savage, R. Zegarac, W. Kovac, C. Hall, J. Chen, R. Last, D. Kramer,
submitted).
One particularly vital parameter is electron transfer rates through photosystem II, or
linear electron flow (LEF). This has traditionally been done by using chlorophyll a fluorescence

144

to calculate the quantum yield of PS II ($II) (187, 228), which is then multiplied by light
-

-2

intensity and leaf absorbance to yield a rate of electron transfer on an area basis (µmoles e m
-1

s ). This method, and the variations of this method, correlate well with O2 evolution and CO2
assimilation, and have become a cornerstone measurement in photosynthesis research. However,
this calculation of LEF makes many assumptions, which may not be valid when applied to the
mutant or ecotype populations that are currently the subject of large-scale imaging experiments.
During large screening programs it may be beneficial to isolate subpopulations or
phenotypes of interest by these more simplistic and rapid methods; however, after isolation of
the desired phenotype, a careful consideration of the elements that determine electron transfer
rates, and appropriate corrections for these may yield even more vital information as to the
nature of any perceived changes. In this work we discuss the variables applied in the traditional
calculation of LEF, the assumptions that are made in the application of these variables, and
conditions in which violation of these assumptions may make a large difference in perceived
electron transfer rates.
A2.2.A What do we need to know to calculate LEF rates?
To measure electron transfer rates through PS II, we need to know several parameters:
The incident light intensity, !, the fraction of the light that is absorbed by PS II bound
chlorophylls, fII, and the quantum yield of PS II, i.e. the fraction of open PS II centers, $II (7,
-

187). These parameters combine to give electron transfer rates on a per leaf area basis (µmoles e
-2 -1

m s ).

145

Eq. A2.1
!"#! ! !! ! !!! ! ! !!!

This equation is similar to past methods (7, 69, 227, 229), with one key difference; the fII
parameter is difficult to directly measure where:

Eq. A2.2
!!!! ! !

!!!!!
!!!!! ! ! !!!!

or, fII is fraction of total chlorophyll bound to PS II. Assuming that all chlorophylls absorb
evenly and all chlorophylls are bound, or chlorophyll that is not attached to a photosystem are
constant or negligible. In the absence of a simple way to determine the fraction of total
chlorophyll bound to PS II, previous corrections for quantifying LEF included measuring total
leaf absorptivity using an integrating sphere, and assuming that all light absorbed was directed
toward photosystems (69).
In the absence of a state transition, or differences in antenna size, it has been generally
accepted to assume total absorbance (per area unit) is attributed to equal absorbance of PS II and
PS I bound chlorophylls and that these are equally distributed between the photosystems.
However, accounting for LEF differences when there is a state transition (even a small one) is
not easily done. There may also be changes in antennae size that do not appear as a state
transition, i.e. if there is not a change in the excitation ratio of the photosystems a state transition
is not perceived. This problem is compounded if there is a combination of these differences when
146

comparing mutants within a species, or plants between species. In these cases, it may be vital to
quantify fII. To assign a value to fII, we can begin by looking at the rate of fluorescence rise with
low illumination, in the presence of DCMU (230).
Under these conditions the rate constant of the rise is dependent on the size of the
antenna, BII , and the quantum efficiency of PS II, $II

Eq. A2.3
!!! ! !!! ! ! ! !!"#$ !!

and which rearranges to:

Eq. A2.4
!!! ! ! !

!!"#$
!!!!!
! !!
!!
!!!
!!"##!

in which BII is proportional to the number of chlorophylls bound to each PS II. Then by
substitution:

Eq. A2.5
!
!!"#$
!!! ! ! !
!
!!
!!

147

Where !!"#$ is the halftime of the fluorescence rise, and !! /!! is the maximal quantum yield of
PS II (7).The incident light intensity would also be factor, however, if the intensity is held as a
constant, it can be ignored.
This tells us the relative amount of chlorophyll attached to each individual PS II, but does
not yet give us the fraction of total chlorophyll that is bound to a PS II. To get at this, we need to
know the concentration of PS II. It is much simpler (and accessible) to measure relative PS I
content by total absorbance changes at 820 nm in DCMU treated leaves where:

Eq. A2.6
!"# ! ! ! !!!!"#!" !

Where:

Eq. A2.7
!!!!"#!" ! ! !!!"#!"!!"#!! ! ! !!!"#!"!!"#$

in which the concentration of PS I is proportional to the difference in absorbance of 820 nm light
of the reduced (dark adapted) and completely oxidized (during the second saturation flash) forms
of P700 (121) (Figure A2.1).
By giving two successive saturating flashes in the presence of DCMU it should be
possible to completely oxidize PS I, as the PC pool is oxidized by PS I following the first flash.
Partially oxidized PS I remains as indicated by the sustained increase of 820 nm absorbance

148

Figure A2.1 Representative experimental data for the calculation of [PSI]. !A820 nm increases
+

with concentration of P700 . First, a 2 second saturation flash is applied to a dark-adapted,
DCMU infiltrated leaf. The following dark interval does not decay to dark-adapted leaves, as
+

DCMU blocks electrons from PS II from reducing P700 . A second 2 second saturating flash is
able to then further (and fully) oxidize the remaining P700. The difference between the !A820 nm
of the second saturating flash and the !A820 nm of the dark-adapted state is proportional to [PSI].

149

during the dark interval. A second flash should completely oxidize PS I, allowing a total
absorbance change to be measured which is relative to total PS I content on a per leaf area basis.
Then, to calculate PS I/PS II, we use the electrochromic shift of the carotenoids at 520
nm in response to an electric field across the thylakoid membrane (Figure A2.2). A short
multiple-turnover saturating flash (10 µs) will excite the photosystems and induce charge
separation and electric field. In the presence of DCMU there is no contribution to electric field
by protons, therefore, the initial amplitude of the ECS is proportional to the charge separation of
both PS I and PS II. PC reduces PS I after oxidation by the first flash, leading to a long-lived
maintenance of electric field. A second flash drives charge separation only through PS I,
allowing the calculation of ECS attributed to PS I alone [discussed in Kramer and Crofts (129)].
This allows us to calculate the fraction or ratio of PS II to PS I, or E:

Eq. A2.8
!! ! !

!"#!" ! !"#!"
!!"##!
!!
!"#!"
!!"#!

And if we take the product of these 3 parameters:

Eq. A2.9
!!! ! !! ! !!!!"#!" ! ! !

!!!!!
!"##
!
! !"# !
!"##
!"#

which simplifies to ChlII, or relative number of chlorophylls bound to PS II. To determine the
fraction of chlorophylls bound to PS II, we need to know the total chlorophyll content per leaf
150

Figure A2.2 Representative experimental data for the calculation of E. Under an electric
field, there is a shift in the absorbance spectrum of the carotenoids towards 520 nm, the so-called
electrochromic shift. A short saturating flash (downward arrows) is applied to a dark-adapted,
DCMU infiltrated leaf and there is a sharp rise in !A520 nm due to charge separation at PS I and
PS II. A second saturation flash generates further ECS due to charge separation at PS I alone,
and E is calculated as described in Eq. A2.8.

151

area. This may be determined as described in Chapter 4, or in the absence of a change in the
composition of the carotenoids to chlorophyll ratios, the !"#!" . However, without this
information, if we use the relative number of chlorophylls bound to PS II for the fII parameter, in
place of the fractional amount of total chlorophyll:

Eq. A2.10
!!!! ! ! !!! ! !! ! !!!!"#!"

Under normal growth chamber conditions, or in mutants that do not display large
differences in either state transitions or chlorophyll a/b, this extensive correction of LEF may not
be necessary. However, under conditions where there is a large state transition, less
pigmentation, or skewed chlorophyll a/b, we have outright violated the assumptions generally
made while calculating LEF, and a very careful correction is required.
This thesis presents several mutants in which it is likely that the standard assumptions
within the LEF calculation are likely violated (Chapters 3 and 4) despite careful corrections
described within those manuscripts. There are also likely multiple instances in the literature that
would likely benefit from a more complete characterization of LEF rates. One such mutant that is
currently being studied in multiple labs is an Arabidopsis mutant with increased rates of cyclic
electron flow around photosystem I (69) (E-M Aro, E. Tyystjärvi, personal communication). It
has been proposed that hcef1 has increased rates of CEF through the NDH complex (69). This
interpretation of the data is dependent on a precise measurement of LEF in this mutant.
+

Traditional methods suggested a greater increase in the slope of vH /LEF than in the slope of

152

+ -

$I/$II (69), which could be interpreted as an increase in the H /e stoichiometry of CEF over the
so-called LEF, and would require the involvement of a proton-pumping plastoquinone reductase.
+ -

To truly attempt a calculation of the H /e ratio of CEF in vivo, it is vital that we develop a more
careful approach to LEF quantification. Therefore, we developed a combined set of methods that
allow us to do this. In the process we have discovered thylakoid architecture changes in hcef1
that have direct implications for CEF levels described in Livingston et al. (69), and functional
properties of NDH.
A2.3 Results and Discussion
A2.3.A PS II antenna size
-2 -1

When leaves infiltrated with DCMU are iluminated at low (>100 µmol photons m s )
light, it allows a slow fluorescence increase, of which the halftime and quantum yield can be
easily calculated. The inverse of the !!"#$ is proportional to the product of the antenna size (BII)
and quantum yield of PS II ($II) (see Eq. A2.3) and simple rearrangement of parameters allows
us to calculate BII (see Eq. A2.4). In Col-0 and hcef1 these values were calculated as 0.76 and
1.26 (from Eq. A2.5) when using 700 nm illumination as necessitated by Chapter 5 conditions,
this indicates an effective increase in hcef1 PS II antennae size of 66% over Col-0.
A2.3.B Relative PS I content
Figure A2.3 shows the absorbance changes of P700 at 820 nm from dark to light in
DCMU treated leaves of Col-0 (Figure A2.3A) and hcef1 (Figure A2.3B). As we show in Eq.
A2.6, the total amplitude of !A820 nm is proportional to PS I content (per area unit). Therefore,

153

Figure A2.3 Decreased [PSI] in hcef1. !A820 nm in dark-adapted, DCMU infiltrated leaves of
Col-0 (A) and hcef1 (B) after two 2 second saturating flashes as described in Figure A2.1.

154

we use this value as [PSI]. We see 58% less [PSI] in hcef1 than in Col-0 (1.4 and 2.4
milliabsorbance units, respectively).
A2.3.C PS II/PS I content
Using Eq. A2.8 and the amplitude of the ECS induced by two multiple turnover flashes
100 ms apart (Figure A2.4), we are able to calculate E in Col-0 and hcef1 (0.67 and 0.4
respectively). This measurement reveals changes in the makeup of the thylakoid membrane of
hcef1. Specifically, hcef1 has less PS II/PS I content. Coupled with the relative PS II antenna size
increase in hcef1, this data may explain the perceived state I condition of hcef1 (D. Strand and D.
Kramer, unpublished), however to fully determine this PS I antenna size must be addressed.
This method allows careful determination of relative LEF values, and may be of vital
importance when comparing LEF rates in not only mutants, but under conditions where changes
in light absorbance may occur, such as different quality of light. In Chapter 5 we use 700 nm
light as an actinic source to poise the system with a more oxidized PS I pool, in which case the
fraction of total photons absorbed by PS II are certainly lower than the fraction absorbed from
625 nm LEDs used in previous studies (33, 69, 72) (Chapters 3 and 4). However, when BII is
calculated from a 625 nm LED and a 700 nm LED, both hcef1 and Col-0 lose the same fraction
of total photons absorbed, suggesting that despite the change in antennae size, there is no
difference in the makeup of the antennae structure of hcef1 (Data not shown). This method of
calculating LEF may also be of value when comparing the effects of different wavelengths on
photosynthesis or photoinhibition, where less absorbance of a specific wavelength may lead to
more open PS II centers, and therefore higher perceived rates of LEF.
While there have been no newly described methods in this manuscript, the combination
of methods allow for a new way to calibrate LEF across a variety of conditions. This does not
155

Figure A2.4 Altered [PSII]/[PSI] in hcef1. !A520 nm in dark-adapted, DCMU infiltrated leaves
of Col-0 (A) and hcef1 (B) after two 10 µs saturating flashes as described in Figure A2.2.

156

correct any assumptions that may be violated within the $II measurement, and any attempt to do
so has been described extensively elsewhere (179, 187, 228, 231, 232).
A2.4 Materials and Methods
A2.4.A Plant materials and growth conditions
Plants and growth conditions are as described in Chapter 5.
A2.4.B In vivo spectroscopy
Chlorophyll a fluorescence and absorbance spectroscopy was performed on a
spectrophotometer described in detail previously (166). In all assays leaves were infiltrated with
50 µM DCMU by incubating detached leaves in the dark for 3 hours between 2 saturated lab
tissues.
A2.5 Acknowledgements
I would like to thank Dr. Nicolas Fisher, Dr. Atsuko Kanazawa, and Dr. Jeff Cruz for
helpful discussions. This work was supported by Grant DE-FG02-11ER16220 from the
Photosynthetic Systems program from Division of Chemical Sciences, Geosciences, and
Biosciences, Office of Basic Energy Sciences of the US Department of Energy (to David M.
Kramer).

157

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