INTERACTIONS OF THE PHOTOSYNTHETIC MACHINERY WITH THE PROTON MOTIVE FORCE: LIMITATIONS AND APPLICATIONS FOR IMPROVEMENT By Geoffry Austin Davis A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Cell and Molecular Biology ÐÐDoctor of Philosophy 2018ABSTRACT INTERACTIONS OF THE PHOTOSYNTHETIC MACHINERY WITH THE PROTON MOTIVE FORCE: LIMITATIONS AND APPLICATIONS FOR IMPROVEMENT By Geoffry Austin Davis The light reactions of photosynthesis convert harvested light into chemical energy that can be utilized by cells for metabolism. Through the translocation of protons across the thylako id membrane coupled to electron transfer reactions, the photosynthetic proton motive force ( pmf ) is used to drive the production of ATP synthesis. Detailed studies have characterized the molecular processes of pmf Ðmediated feedback regulation of photosynth esis via changes in luminal pH from the light -induced proton gradient ( ! pH) . While it is now well established that the photosynthetic pmf in higher plants consists of both a ! pH and an electric potential ( !!), the impact that !! exerts on photosynthesis in vivo is mostly unstudied. The !! component, howeve r, influences the relative free -energy between redox mediators of electron transfer within membrane complexes . We found that in plants, a large in vivo !! increases photoinhibition through photosystem II (PSII) damage. High !! levels were observed in mutants with high steady -state pmf levels, as well as in wild type plants during light fluctuations. The increase in photoinhibition is primarily due to increased yields of electron recombination in PSII, whic h generate reactive oxygen species (ROS). The yield of PSII recombination when !! is large is mediated by ! pH-dependent photosynthetic downregulation to decrease the concentration of reduced electron acceptors in PSII (Q A-) capable of recombining and generating ROS. The ability to regulate photosynthetic light capture and electron transfer via pH Ðdependent processes as well as the need for photosynthetic organisms to mitigate a large !! leads me to propose that the photosynthetic organisms have evolve d regulatory processes to mediate the bioenergetic limitations imposed by the !! effect on electron transfer. In a population of natural Arabidopsis thaliana accessions, variation in the kinetics of activating and deactivating pH -dependent downregulation o f photosynthesis allowed the genetic loci responsible for the kinetics within this population to be mapped. These results suggest that photosynthetic organisms have evolved multiple mechanisms to regulate how rapidly photosynthesis is regulated by the pmf . The partitioning of pmf between !! and !pH allows a plant to balance the induction and relaxation of photoprotective mechanisms at the detriment of light utilization, while minimizing the impact of !!-mediated PSII recombination and ROS production. The pr ocesses wor k in concert to minimize potential photo damage and loss of productivity that occurs due to biophysical alterations in electron transfer processes mediated by !!. Copyright by GEOFFRY A. DAVIS 2018 iv ACKNOWLEDGEMENTS My dissertation research has been tremendously impacted by the involvement of the Kramer lab as a whole, who have fostered a friendly, non -competitive environment full of creativity and, at times, weirdness. My mentor, Dr. David Kramer, has allowed me to grow as a scientist both in experimental work as well as communicating science. While at times challenging, I have greatly appreciated the determination for excellent science he pushes me towards. I have had the pleasure to work on many projects with Dr. Mio Satoh -Cruz, from whom I have benefitted and learned her great organizational skills. Dr. Nicholas Fisher provides a wealth of knowledge for in vitro enzymology as well as all things tea and cricket. With Dr. Fisher, I had the pleasure of working with a former graduate student and postdoctoral researcher Dr. Deserah Strand on cyclic electron flow projects. The cyclic team was a great introduction to the Kramer lab during my rotation, and the energy and passion Dr. Strand has for researc h is inspiring. Much of my work would not have been possible without the technical assistance of Dr. Jeff Cruz, Robert Zegarac, and Nathan Galbreath. The engineering assistance they have provided has allowed me to take instruments up to 11 for my experime nts. Linda Savage and David Hall, along with the undergraduate assistants they have employed, have been critical assistants in my many plant imaging experiments as well as resources for growing and managing Arabidopsis. Although my work did not often lea d me into the algae group, I have appreciated the interest and feedback on experiments from Dr. Ben Lucker. v Many experiments were carried out in a timely and organized manner thanks in large part to the excellent assistance of Abigail Crampton while work ing as an undergraduate research assistant. I have had the opportunity to conduct my dissertation research in the Michigan State University Department of Energy Plant Research Laboratory. The resources, expertise, and openness of the different lab groups has been an outstanding working environment. In particular, I have appreciated the opportunities to interact with Dr. John Froehlich, and learn from members of Dr. Tom SharkeyÕs carbon assimilation lab and Dr. Daniel DucatÕs photosynthetic synthetic biolog y lab. Within my graduate department, I have enjoyed the weekly research seminars and the interactions with Dr. Sue Conrad and Dr. Kathy Meek. Outside of formal department affiliations, the interactions between labs at Michigan State University has allowed me to comfortably discuss questions with experts in many fields as they have occurred, and I am grateful for the openness that has been fostered. Lastly, I would like to thank my partner Siobhan Cusack, who has worked with me as we have both navigated o ur dissertations. Your optimism and support have helped with the many difficulties graduate student life provides, and I think that after five years you now know what the proton motive force does. vi TABLE OF CONTENTS LIST OF TABLES ......................................................................................................................... ix! LIST OF FIGURES ........................................................................................................................ x! KEY TO ABBREVIATIONS ..................................................................................................... xiii i! Chapter 1 ......................................................................................................................................... 1!Regulation of photosynthesis in higher plants by the thylakoid proton motive force .................... 1!1.1 Photosynthetic electron tr ansfer is tightly coupled to the generation of a trans -thylakoid proton motive force. .................................................................................................................... 2!1.2 The photosynthetic pmf ! pH is a key feedback regulatory loop. ......................................... 3!1.3 Trans Ðthylakoid !! regulation is poorly un derstood but is not without consequences to photosynthesis. ............................................................................................................................ 6!1.4 The ATP synthase cÐring stoichiometry dictates the efficiency of pmf -generated ATP synthesis. ..................................................................................................................................... 7!1.5 A large ATP synthase c-ring stoichiometry may help stabilize pmf partitioning into ! pH to minimize !!. .............................................................................................................................. 9!1.6 Photosynthetic regulation limits the influence of !! on electron transfer. ........................ 10! Chapter 2 ....................................................................................................................................... 13!Limitations to photosynthesis by proton motive force -induced photosystem II photodamage 1 ... 13!2.1 Abstract ............................................................................................................................... 14!2.2 Introduction ......................................................................................................................... 15!2.3 Results ................................................................................................................................. 17!2.4 Discussion ........................................................................................................................... 36!2.5 Materials and Methods ........................................................................................................ 40!2.5.1 Plant materials and growth conditions. ........................................................................ 40!2.5.2 Generation of chloroplast ATP synthase "-subunit minira (minimum recapitulati on of ATPC2) mutants. .................................................................................................................. 40!2.5.3 Isolation of tightly coupled chloroplasts and intact thylakoids. .................................. 43!2.5.4 Spectroscopic measurements. ...................................................................................... 43!2.5.5 Estimation of recombination rate. ................................................................................ 45!2.5.6 Chlorophyll fluorescence imaging. .............................................................................. 46!2.5.7 Photoinhibition of detached leaves. ............................................................................. 47!2.5.8 Protein analysis. ........................................................................................................... 48!2.5.9 1O2 detection. ............................................................................................................... 48!2.5.10 Data analysis. ............................................................................................................. 49!2.6 Acknowledgements. ............................................................................................................ 49! Chapter 3 ....................................................................................................................................... 83!Hacking the thylakoid proton motive force for improved photosynthesis: modulating ion flux rates that control proton motive force partitioning into !! and ! pH ........................................... 83!vii 3.1 Abstract ............................................................................................................................... 84!3.2 Introduction ......................................................................................................................... 85!3.3 Interactions of the pmf with PSII: Importance for energy storage a nd photodamage. ....... 88!3.4 Computational exploration of the effects of altering pmf storage kinetics on q E and field recombination Ðinduced photodamage. ..................................................................................... 90!3.5 The effects of ! pH and !! on PSII recombination a nd 1O2 production in vivo . ............... 91!3.6 Control of the extent and kinetics of pmf partitioning. ....................................................... 94!3.7 Can we improve photosynthesis by modifying pmf partitioning to make photosynthesis more robust? ............................................................................................................................ 102!3.8 Acknowledgements. .......................................................................................................... 107! Chapter 4 ..................................................................................................................................... 111!The electric field component of the photosynthetic proton motive force increases the yield of PSII recombination in vivo during light fluctuations .................................................................. 111!4.1 Abstract ................................................................................................................................. 112!4.2 Introduction ....................................................................................................................... 113!4.3 Materials and Methods ...................................................................................................... 118!4.3.1 Plant and growth conditions. ...................................................................................... 118!4.3.2 Spectroscopic measurements of delayed fluorescence, prompt fluorescence, and the electrochromic shift. ........................................................................................................... 118!4.3.3 Determination of delayed fluorescence yields. .......................................................... 119!4.4 Results ............................................................................................................................... 120!4.4.1 Abrupt increases in light intensity increase the yield of delayed fluorescence. ........ 120!4.4.2 Alterations in the regulation of PSII Q A redox state leads to changes in field -induced increases in recombination. ................................................................................................. 122!4.5 Discussion ......................................................................................................................... 124!4.6 Acknowledgements ........................................................................................................... 129! Chapter 5 ..................................................................................................................................... 130!Natural variation in the activation and dissipation of nonphotochemical quenching in Arabidopsis thaliana subjected to fluctuating light .................................................................... 130!5.1 Abstract ............................................................................................................................. 131!5.2 Introduction ....................................................................................................................... 132!5.3 Materials and Methods ...................................................................................................... 135!5.3.1 Plant materials and growth conditions ........................................................................... 135!5.3.2 Chlorophyll fluorescence imaging ................................................................................. 135!5.3.3 QTL analysis .............................................................................................................. 136!5.3.4 Protein extraction and western blotting ..................................................................... 137!5.4 Results ............................................................................................................................... 137!5.4.1 Variation in NPQ kinetics between Arabidopsis accessions ..................................... 137!5.4.2 QTL mapping of NPQ kinetics .................................................................................. 140!5.4.3 QTL mapping of rapid responses to light fluctuations .............................................. 142!5.5 Discussion ......................................................................................................................... 147!5.6 Acknowledgements ........................................................................................................... 151! viii Chapter 6 ..................................................................................................................................... 158!Conclusions and future directions ............................................................................................... 158!6.1 Abstract. ............................................................................................................................ 159!6.2 Conclusions. ...................................................................................................................... 159!6.3 Future directions. .............................................................................................................. 160! APPENDIX ................................................................................................................................. 163 REFERENCES ........................................................................................................................... 174!ix LIST OF TABLES Table 2.1: Oligonucleotide sequences utilized for adapter ligation mutagenesis. ........................ 71! Table 2.2 Oligonucleotide sequences utilized for adapter ligation mutagenesis to introduce secondary mutations. ..................................................................................................................... 72! Table 2.3: Oligonucleotide sequences utilized for splicing by overlap extension PCR. .............. 73! Table 2.4: Synthetic gene constructs incorporating mul tiple ATPC2 mutations into ATPC1. .... 74! Table 2.5: Timing and light profile of imaging day 1 ................................................................... 75! Table 2.6: Timing and light profile of imaging day 2 ................................................................... 76! Table 2.7: Timing and light profile of i maging day 3 ................................................................... 78! Table 2.8: Chlorophyll content of wild type (Ws -2) and minira leaves. ...................................... 82! Table 5.1: List of QTLs identified for time -dependent NPQ responses to fluctuating light. ..... 152! Table 5.2: List of QTLs identified for time -dependent changes in "f amplitude. ....................... 154! x LIST OF FIGURES Figure 1.1: Electron and proton transfer steps of the photosynthetic light reactions. .................... 5! Figure 2.1: "-subunit mutations alter photosynthetic proton efflux. ............................................. 18! Figure 2.2: Dynamic ligh t conditions enhance pmf dependent phenotypes. ................................. 21! Figure 2.3: Elevated pmf leads to PSII photodamage. .................................................................. 23! Figure 2.4: The dependence of photoinhibition on the redox state of Q A. ................................... 25! Figure 2.5: Photoinhibition is strongly correlated with !! but not ! pH in minira lines. ............ 26! Figu re 2.6: In vitro manipulation of the !! alters PSII S 2QA- recombination rates. .................... 28! Figure 2.7: Induction of !! and 1O2 production under fluctuating light in wild t ype plants. ...... 30! Figure 2.8: Schemes for the trans -thylakoid !!-induced acceleration of recombination reactions in PSII and subsequent production of 1O2. .................................................................................... 35! Supplemental Figure 2.1: Whole plant fluorescence imaging phenotyping of minira 3Ð1 mutant. ....................................................................................................................................................... 50! Supplemental Figure 2.2: Increased pH -dependent quenching correlates with increased photoinhibitory quenching. ........................................................................................................... 51! Supplemental Figure 2.3: Whole plant fluorescence imaging phenotyping of minira 11Ð1 mutant. ....................................................................................................................................................... 52! Supplemental Figure 2.4: Whole plant fluorescence imaging phenotyping of minira 14Ð1 mutant. ....................................................................................................................................................... 53! Supplemental Figure 2.5: Whole plant fluorescence imaging phenot yping of minira 12Ð2 mutant. ....................................................................................................................................................... 54! Supplemental Figure 2.6: Whole plant fluorescence imaging phenotyping of minira 8Ð1 mutant. ....................................................................................................................................................... 55! Supplemental Figure 2.7: Whole plant fluorescence imaging phenotyping of minira 6Ð2 mutant. ....................................................................................................................................................... 56! xi Supple mental Figure 2.8: Whole plant fluorescence imaging phenotyping of minira 4Ð2 mutant. ....................................................................................................................................................... 57!Supplemental Figure 2.9: Whole plant fluorescence imaging phenotyping of minira 6Ð1 mutant. ....................................................................................................................................................... 58! Supplemental Figure 2.10: Whole plant fluorescence imaging phenotyping of minira 7Ð1 mutant. ....................................................................................................................................................... 59! Supplemental Figure 2.11: Whole plant fluorescence imaging phenotyping of minira 3Ð2 mutant. ....................................................................................................................................................... 60! Supplemental Fig ure 2.12: Whole plant fluorescence imaging phenotyping of minira 4Ð1 mutant. ....................................................................................................................................................... 61! Supplemental Figure 2.13: Whole plant fluorescence imaging phenotyping of minira 9Ð1 mutant. ....................................................................................................................................................... 62! Supplemental Figure 2.14: Whole plant fluorescence imaging phenotyping of minira 4Ð3 mutant. ....................................................................................................................................................... 63! Supplemental Figure 2.15: Whole plant fluorescence imaging phenotyping of minira 12Ð3 mutant. .......................................................................................................................................... 64! Supplemental Figure 2.16: Whole plant fluorescence imaging phenotyping of minira 2Ð2 mutant. ....................................................................................................................................................... 65! Supplemental Figure 2.17: Reduction kinetics of P700 +. ............................................................. 66! Supplemental Figure 2.18: The electric field component of the pmf dominates under high pmf conditions. ..................................................................................................................................... 67! Supplemental Figure 2.19: Tobacco ATPC1 antisense knockdown increase !! partitioning under high pmf conditions. ...................................................................................................................... 68! Supplemental Figure 2.20: Uncoupling !! decreases SOSG fluorescence in minira 3Ð1. ......... 69! Supplemental Figure 2.21: Fluctuations in light intensity result in transient ECS spikes. ........... 70! Figure 3.1: Photosynthetic electron transfer energetics are influenced by membrane orientation and membrane potential. ............................................................................................................... 92! Figure 3.2: Illustration of the factors contri buting to the balancing of pmf into !! and ! pH during different phases of illumination. ........................................................................................ 95! Figure 3.3: Simulated responses of thylakoid pmf components, linear electron flow, ion fluxes and 1O2 production during a 5 -min light pulse. ............................................................................ 98!xii Figure 3.4: Simulated responses of thylakoid pmf components, linear electron flow (LEF) and 1O2 production during a 1 -h light sine wave. ....................................................................... 102! Figure 3.5: Effects of thylakoid counter -ion fluxes on the ATP/NADPH budget of photosynthesis. ............................................................................................................................ 104! Supplemental Figure 3.1: Simulatio ns of the amplitudes of pmf parameters induced by a single -turnover flash hitting all photosystem II (PSII), but no photosystem I (PSI). ............................ 108! Supplemental Figure 3.2: Simulated responses of thylakoid pmf components, linear electron flow, ion fluxes, and 1O2 production during a 5 -min light pulse. .............................................. 109! Supplemental Figure 3.3: Simulated responses of thylakoid pmf components, linear electron flow, and 1O2 production during illumination with a 1 -hour sine or square waves. ................... 110! Figure 4.1: Photosystem II electron transfer reactions successively increase the distance of transmembrane charge separation. .............................................................................................. 116! Figure 4.2: Fluctuating light increases PSII recombination in vivo . ........................................... 122! Figure 4.3: Deficiencies in pH -dependent NPQ antenna quenching increase #DF due to increased PSII center with Q A-. ................................................................................................... 124! Figure 5.1: The 19 founding parental accessions of the Arabidopsis MAGIC population display a range of NPQ induction and dissipation kinetics during fluctuating light. ................................. 138! Figure 5.2: Differential content of proteins mediating the rapidly reversible pH -dependent q E component of NPQ Arabidopsis accessions. .............................................................................. 140! Figure 5.3: Kinetics of NPQ responses in the Arabidopsis MAGIC population under fluctuating light. ............................................................................................................................................ 141! Figu re 5.4: Time -dependent identification of NPQ QTL during fluctuating light in Arabidopsis MAGIC lines. .............................................................................................................................. 142! Figure 5.5: The kinetics and amplitude of chlorophyll fluorescence quenching varies among the Arabidopsis MAGIC founders and MAGIC population during light fluctuations. .................... 145! Figure 5.6: Time -dependent changes in the amplitude of "f quenching during light fluctuations. ..................................................................................................................................................... 147!xiii KEY TO ABBREVIATIONS 1O2 Singlet oxygen !GATP Free -energy of hydrolysis of ATP ! pH pH gradient !! Electric field #II Quantum yield of photosystem II activity in the light #DF Quantum yield of delayed fluorescence "f Fluorescence yield ATP Adenosine tri -phosphate ADP Adenosine di -phosphate ANOVA Analysis of variance DCMU 3-(3,4-dichlorophenyl) -1,1-dimethylurea DF Delayed fluorescence e- Electron ECS Electrochromic shift ECS inv Inversion of the electrochromic shift ECS ss Electrochromic shift decay steady -state ECS t Total amplitude of the electrochromic shift Fo Minimum chlorophyll fluorescence from dark -adapted sample FRIP Field recombination -induced photodamage Fs Fluorescence emission from sample in the light FV/FM Maximum quantum efficiency of photosystem II xiv gH+ Proto n conductivity of the ATP synthase H+ Proton K+ Potassium cation LEF Linear electron flow MAGIC Multi-parent advanced generation inter -cross NPQ Nonphotochemical quenching NADP+/NADPH Oxidized/reduced nicotinamide adenine dinucleotide phosphate ODE Ordinary differential equation OEC Oxygen -evolving complex P Primary reaction center donor P680 Primary photosystem II chlorophyll electron donor P700 Primary photosystem I chlorophyll electron donor 1P Singlet electron state of primary reaction center donor 3P Triplet electron state of primary reaction center donor P* Excited state of primary reaction center donor PAR Photosynthetically active radiation PC Plastocyanin Pheo Pheophytin Pi Inorganic phosphate pmf Proton motive force PMT Photomultiplier tube PQ Plastoquinone PQH2 Plastoquinol xv PSI Photosystem I PSII Photosystem II QA Photosystem II primary electron acceptor quinone QB Photosystem II secondary electron acceptor quinone qE Energy -dependent chlorophyll a fluorescence quenching qI Photoinhibitory chlorophyll a fluorescence quenching qL Fraction of photosystem II with Q A QTL Quantitative trait locus ROS Reactive oxygen species S1/ S2/ S3/ S0/ Sn S-state of the oxygen -evolving complex SNP Single nucleotide polymorphism SOSG Singlet oxygen sensor green VDE Violaxanthin de -epoxidase YZ Photosystem II D1 protein tyrosine Z ZEP Zeaxanthin epoxidase 1 Chapter 1 Regulation of photosynthesis in higher plants by the thylakoid proton motive force Geoffry A. Davis 2 1.1 Photosynthetic electron transfer is tightly coupled to the generation of a trans -thylakoid proton motive force. Conversion of light into chemical energy in oxygenic photosynthetic membranes involves the cap ture of light and energy funnel ing through a series of redox intermediates to ultimately generate NADPH from NADP +. These electron transfer steps are tightly coupled to proton transfer reactions across the membrane, stabilizing th e electro n transfer reactions via charge -coupled movement and generating a potential energy store in the form of the proton motive force (pmf ) (1). Through the activity of the pmf , ATP synthesis via the rotary catalysis of the ATP synthase generates ATP fo r metabolic processes required for cellular proliferation. The pmf is composed of both a proton gradient ( ! pH) and an electrochemical gradient ( !!) created via redistribution of H + and other ion species between the thylakoid lumen and chloroplast stroma by photosynthetic processes (2). Both components are thermodynamically equivalent in driving the ATP synthase (3) and the total driving force for ATP synthesis can be described as: !"# !!!!!!!!!!!!"!!!"!!! (Eq. 1) where !!i-o and ! pHo-i represent the electric field and proton gradient calculated as the difference in concentrations between the inside (lumen) and outside (stroma) , R is the universal gas constant, and F is FaradayÕs constant. While both components of the pmf are thermodynamically equivalent drivers of ATP synthesis (3, 4), as can be seen from Eq. 1, !! and ! pH are not kinetically equivalent, with a larger ! pH requi red to reach an equivalent !! driving force. The pmf is primarily dissipated by H+ efflux from lumen through the ATP synthase, but both pmf generation and dissipation are tightly regulated during photosynthesis, requiring coordination of the electron transfer processes with substrate acceptor availability in the chloroplast stroma (e.g. 3 NADP+, ADP, P i) to match the rapid processes of electron and proton transfer to the slower metabolic processes occurring in cells. 1.2 The photosynthetic pmf !pH is a key feedback regulatory loop. During the course of oxygenic photosynthesis, the vectorial movement of electrons through membrane embedded proteins is tightly coupled to the deposition of protons into the thylakoid lumen (Fig. 1 .1). This movement of prot ons both stabilizes the transfer of the electrons across the membrane and acts as a chemical store of potential energy through the generation of a proton gradient. While the pmf is dissipated through the ATP synthase to generate ATP, acidification of the t hylakoid lumen during ! pH generation acts as a feedback signal to various components of the light reactions (reviewed in 5). As the ! pH increases, protonation of violaxanthin deepoxidase occurs as the lumen pH decreases below ~6.5 and is fully active at pH ~5.8, leading to the accumulation of the carotenoid derivative zeaxanthin in the thylakoid membrane, which has a photoprotective role and is part of the energy -dependent nonphotochemical quenching (q E) processes (Fig. 1.1) (6). Similarly, protonation of lumen -exposed glutamate residues of the peripheral antennae protein PsbS also contributes to q E (7), which, in combination with zeaxanthin accumulation, contributes to the total q E response (8). Following protonation of PsbS and accumulation of zeaxanthin , excitation energy is quenched in the antenna pigment protein complexes via energy transfer from singlet or triplet excited state chlorophylls to closely localized antenna carotenoid and zeaxanthin (9). Acidification of the thylakoid lumen also acts as a thermodynamic limitation to the proton releasing steps of photosynthesis. Oxidation of plastoquinol by cytochrome b6f results in the release of two protons into the lumen per plastoquinol oxidized, which is coupled to the 4 continuous activity of the Q -cycl e, resulting in three protons transferred from the stroma into the thylakoid lumen per electron from PSII (10). As the lumen acidifies, the rate of plastoquinol oxidation at the cytochrome b6f Qo site decreases tenÐfold from pH 7.5 to pH 5.5 (11, 12), limiting the rate of photosynthetic electron transfer (reviewed in (13, 14). Similarly, proton release from the PSII oxygen evolving complex (OEC) has also been shown to be kinetically limited by acidification of the lumen (15). Reduction of the primary PSII e lectron donor (P 680+) is facilitated by redox intermediate states (S -states) of the Mn 4Ca OEC complex, during which changes in Mn oxidation status due to electron transfer to reduce P 680+ are couple d to proton deposition in the lumen and water splitting (16, 17). Advancement of the of the OEC S -states become s kinetically limited by the lumen pH in vitro , with the most alkaline p Ka (~4.6) associated with the S 3 to S 0 transition (15). Although a lumen pH of 4.6 is likely far below the physiological lumen pH limit (13), the pH limitation of water splitting would limit the amount of pmf that could be stored as ! pH (15). This pH Ðmediated kinetic control allows the pmf to act as a feedback mechanism to the light capturing and electron transfer components of the thylakoid membrane to tune electron transfer processes to meet the capability of the thylakoid ATP synthase to generate ATP and relieve ! pH, which is dependent upon substrate availability from metabolic processes (18, 19). 5 Figure 1.1: Electron and pr oton transfer steps of the photosynthetic light reactions. Schematic representation of the electron (black dashed lines) and proton (red sold lines) transfer reactions occurring across the thylakoid membrane during photosynthetic linear electron flow. Foll owing absorption of light by antenna pigment protein complexes, excitation transfer and charge separation within photosystem (PS) II and I generates the oxidizing potential capable of splitting water at PSII. Electron transfer from the PSII primary donor P 680 occurs vectorially across the membrane spanning protein through a pheophytin (Pheo) intermediate, the bound primary quinone Q A, to the mobile quinone Q B. P680+ is reduced by electron transfer from the Mn4Ca oxygen -evolving complex (OEC) through a redox active tyrosine (Y Z). Following two successive PSII turnovers, the doubly reduced quinone bound at Q B forms the quinol by protonation and is release into the thylakoid membrane, followed by binding of a new quinone at QB from the quinone pool (PQ). Two mo re PSII turnovers are required to complete the oxidation of water by the OEC and deposition of protons into the lumen. Oxidation of plastoquinol by cytochrome b6f reduces plastocyanin (PC) and translocates more protons into the lumen. Excitation and charge -separation in PSI allow electron transfer from PSI to ferredoxin (Fd) and re-reduction of PSI by PC. Fd is used to generate NADPH from NADP + via ferredoxin -NADP+ reductase (FNR). The proton motive force generated by proton deposition is used to move proto ns from the lumen to the stroma through the ATP synthase c-subunit ring to generate ATP. Lumen acidification regulates multiple thylakoid proteins (blue boxes). Activation of pH -dependent nonphotochemical quenching occurs via protonation of violaxanthin de -epoxidase (VDE) and the conversion of violaxanthin (V) to zeaxanthin and antheraxanthin (Z + A), as well as protonation of PsbS (orange). NADPH -SH -SH ADP + P i ATP H+ Stroma Lumen PQ 2H2O 4H+ + O2 Fd FNR PC Mn4Ca QA YZ Pheo P680 H+ PsbS VDE V Z + A ZE H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ H+ QB b6f H+ H+ PSI PSII 6 1.3 Trans Ðthylakoid !! regulation is poorly understood but is not without consequences to photosynthesis. While the regulatory mechanisms o f ! pH generation have been well -characterized in vitro and in vivo , !!Ðmediated feedback on photosynthesis has been poorly studied. Due to a low electrical capacitance of the thylakoid membrane, generation of !! can occur rapidly from a dark -adapted (fully oxidized) state via the charge disequilibrium that occurs following charge separation within PSII and PSI complexes, producing anion electron acceptors closer to the stromal face and cation holes near the luminal edge within both complexes (20). This !! is rapidly replaced by the redistribution of ions between the stroma (or cytosol in cyanobacteria) and thylakoid lumen (21-25). Although the movement of ions across the thylakoid has been characterized for decades (26), how or if !! regulates photosynthesis in vivo has only recently emerged as a focus of study (2). In Arabidopsis thaliana , the use of knockout or knock down mutants in putative chloroplast ion transporters has identified proteins that , when absent , alter the ! pH/ !! rat io in vivo (27-32), reviewed in (33-35). In mutants with a decreased ! pH/ !! ratio (tpk3 , kea1/kea2 , clce, vccn1, pam71 ), the kinetics of NPQ induction is delayed due to an inability to rapidly collapse !! and generate ! pH as quickly as wild type plants (27, 29-31). This also led to a measured increase in PSII photoinhibition in tpk3 , attributed to the decreased ! pH failing to properly activate q E and preventing proper photoprotection (27). However, recent work with Arabidopsis ATP synthase activity mutants, in which only the buildup of total pmf was purposefully altered but other pmf regulatory proteins remained genetically unchanged, an alteration of the ! pH/ !! ratio was similarly seen unde r high total pmf conditions, suggesting in vivo regulation of pmf partitioning not previously identified that prevents over -acidification of the thylakoid lumen (13). Unlike ion transporter mutants, the 7 activation of NPQ through ! pH generation is not impai red via constitutive mis -regulation of pmf partitioning, but PSII photoinhibition was still prominent when the ! pH/ !! ratio was low (36). This was shown mechanistically to be d ue to destabilization of charge -separated states during vectorial electron trans fer, which decreases the free energy barrier for charge recombination, leading to an increase in singlet oxygen ( 1O2) production via PSII recombination when !! is large (37, 38). While this phenomenon was found during the steady Ðstate in high pmf mutants, the same phenomenon was observed in wild type plants during light fluctuations similar to natural conditions, when a large, transient !! is present following the increase in excitation energy. These findings highlight the importance of !! not only for prov iding pmf to properly balance ATP synthesis (39), but also an emerging role for !!Ðmediated regulation of electron transfer events in vivo . As the core electron transfer protein complexes of photosynthesis ( PSII, cytochrome b6f, and PSI ) are highly conserved across all oxygenic photosynthetic organisms, !! regulation of photosynthetic electron transfer, though poorly studied, is likely to be a prevalent phenomenon occurring under natural (non Ðstatic) conditions with substantial consequenc es for photosynthetic productivity. 1.4 The ATP synthase cÐring stoichiometry dictates the efficiency of pmf -generated ATP synthesis. The dissipation of pmf through the rotary -coupled F 0F1 ATP synthase is utilized in bioenergetic membranes throughout the tree of life, highlighting not only the utility of pmf Ðdriven phosphorylation but also the highly conserved mechanism of ATP synthase activity (reviewed in 40). Rotational movement of the m embrane embedded F 0 portion is driven by a single proton binding residue on each cÐsubunit, which is coupled to the catalytic turnover of the $3%3 hexamer to release 3 ATP molecules per full turnover. Therefore, full turnover of the F 1 8 enzyme is coupled to complete 360¡ rotation of the cÐring, generating 3 ATP per c-subunits. This rotational catalysis mechanism is conserved across species and membranes of various bioenergetics processes, however, the subunit stoichiometry of the cÐring has been shown to var y from 8 Ð15 subunits, although maintaining a single stoichiometry within a species, in turn leading to a wide range of H +/ATP ratios and thus bioenergetic limitations between different species and various cellular compartments membranes. The bioenergetic implications of cÐring stoichiometry have direct consequences on the amount pmf required to drive ATP synthesis (41, 42). While !pH and !! are thermodynamically equivalent drivers of ATP synthesis (Eq. 1), they are not kinetically equivalent. Using ATP synthases with different cÐring stoichiometries, the threshold of pmf activation for ATP synthesis has been shown to vary with the number of c -subunits, with larger values becoming active at lower potentials (43) and having higher activity at lower potential (44). This has been postulated to be due to a decrease in the total rotation required for each cÐsubunit step Ðwise movement as the stoichiometry increases, suggesting that larger cÐring stoichiometries allow those complexes to operate at a lower pmf . Remarkably, the chloroplast ATP synthase cÐring stoichiometry is on the high end of ATP synthase complexes with 14 c-subunits (42, 45). Intriguingly, using published data to derive parameters for ATP synthesis, Silverstein (41) found that even at a nearl y identical pmf , the E. coli and bovine mitochondrial ATP synthases ( c-subunits=10 and 8, respectively) outperform the chloroplast ATP synthase in terms of efficiency by ~25%. This inefficiency specific to the chloroplast ATP synthase was hypothesized to b e due to the increase in pmf storage as !pH component in vivo relative to other membranes, as well as the lower !! threshold for activation (43, 46). While both of these issues may address mathematically why the chloroplast complex is less efficient, they fail to consider 9 why chloroplasts evolved to utilize a higher !pH and lower !!. As described above and thoroughly discussed elsewhere (5, 47), acidifying the thylakoid lumen leads to pH Ðmediated downregulation of photosynthesis, while in other cellular com partments acidification may lead to enzyme damage, or be difficult to acidify due to a larger volume than the thylakoid lumen (48, 49). Chloroplasts, however, still do not solely maintain a !pH as their pmf source, but maintain a balance with !!, likely to avoid deleterio us acid Ðinduced damage to lumen -exposed proteins and protein residues (13, 36). 1.5 A large ATP synthase c-ring stoichiometry may help stabilize pmf partitioning into !pH to minimize !!. If chloroplasts have evolved to supplement a !pH gradient with !! to avoid over Ðacidification, one could speculate as to why chloroplasts d o not maintain a large !! while supplementing with a !pH gradient to activate q E. A larger !! gradient would increase thermodynamic efficiency of chloroplast ATP synthase, but as noted above in chloroplasts, and likely cyanobacteria , due to the highly conserved core electron transfer proteins, a large !! decreases the free energy barriers for electron recombination, leading to increased 1O2 production at PSII (36). Could photosynthetic ATP synthases have evolved to specifically mitigate this !! problem? It is intriguing that the number of c-subunits in photosynthetic organisms analyzed to date are all on the high end ( 13Ð15 subunits) of the determined cÐring stoichi ometries (45, 50, 51). The energy required to catalyze the synthesis of ATP ( ! GATP ) is given by: !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!"#!!!!!!! (Eq. 2) where n is the H +/ATP ratio required to generate each molecule of AT P dictated by the number of cÐsubunits. The high H +/ATP ratio in chloroplasts ( n=4.3) decreases the pmf required to 10 overcome ! GATP , allowing photosynthesis to produce ATP at a lower relative pmf (52), reducing the requirement to maintain a either a large !pH or !! during steady Ðstate photosynthesis. The implications of large cÐring stoichiometry are evident if one considers the energetic consequences that decreasing the stoichiometry would have on photosynthesis. Assuming a !GATP of 40 kJ/mol (53), a decrease in the number of cÐsubunits from 15 to 9 would result in an increase in the pmf required to maintain ! GATP equilibrium from ~83 mV to ~138 mV. If the energy is equally partitioned between !! and ! pH, this leads to an equilibrium ! pH of 0.7 uni ts with a c15 stoichiometry and ! pH of 1.2 units at c9, enough to already begin activating q E processes in the dark (39). Similarly, the !! required just to maintain ! GATP equilibrium increases by nearly 30 mV. Further pmf generation during photosynthesis would therefore exacerbate these increases. Under this hypothetical smaller c9 operating structure, photosynthetic pmf would either need to be limited to a lower total pmf capacity than its current c14 state, or require a dramatic shift in the partitioning into !! to avoid near immediate over -acidification of the thylakoid lumen below ~5.5 ( ! pH 2.6 units assuming stromal pH 7.8) (13, 39). As discussed above, the physical properties of the thylakoid membrane dictate the temporal partitioning of !! and ! pH ge neration. A shift in partitioning in favo r of !! could occur via genetic regulation of counter -ion movement through ion transport expression, or an increase in the buffering capacity of the lumen, which would require massive remodelling of thylakoids or the use of high concentrations of mobile buffering groups such as polyamines (54). 1.6 Photosynthetic regulation limits the influence of !! on electron transfer. A shift of pmf partitioning into more !!, however, will greatly influence the recombination fr equency between charge -separated states in PSII. The recombination rate in PSII depends on the energetics of electron sharing between redox intermediates and changes 11 exponentially with changes in !!. For the charge -separated state(s) forming P +QA-, where P + is the primary electron donor and Q A the non -mobile PSII quinone, these changes correspond to : !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!"#$%&'()*'$( !!!!!!!!!!!!!!!!!!"!!!!"#$ !!!"! (Eq. 3) where [S 2QA- + S3QA-] represents the fraction of PSII containing donor and acceptor side states capable of recombining from Q A-, kr is the intrinsic rate of recombination from S 2/S3QA-, and !Estab is the stabilization free energy of the charge separated state S2/S3QA-, expressed in eV (36). As !! increas es, the ! Estab of charge separated states decreases (37, 55), leading to an increase in the rate of recombination. Therefore, even with relatively small changes in the amount of energy stored as !!, the velocity of recombination will increase dramatically. This likely limits the amount of energy that can be stored safely across photosynthetic membranes as !!, as electron recombination through back Ðreactions can lead to generation of reactive oxygen species (ROS) (36, 56). It therefore appears that a large cÐring stoichiometry, although energetically inefficient (41) may in fact be far more physiologically efficient for organismal survival in photosynthetic membranes. Additionally, the decrease in the chloroplast ATP synthase pmf activation threshold may hav e led to the additional advantage of photosynthetic ATP generation under even low light conditions (46). The increased storage of pmf as ! pH in photosynthetic organisms activates multiple regulatory processes that feedback into the amount of energy held b y QA-. As ! pH builds and q E is activated, excitation quenching in the antenna decreases the fraction of absorbed light transferred to PSII, decreasing the PSII excitation pressure to limit over -reduction and buildup of QA-, which can recombine to generate ROS. Concurrently, downregulation of b6f turnover regulates the redox state of the plastoquinone pool, balancing the PSII excitation pressure with 12 the availability of electron donors. Although pH -mediated activation of NPQ has been well characterized in mu ltiple eukaryotic organisms (57), NPQ mechanisms vary between different photosynthetic lineages, and are not pH regulated in cyanobacteria (58). Comparatively, !! influences on electron transfer are likely to be experienced in all photosynthetic lineages, as the core electron transfer proteins are highly conserved from cyanobacteria to higher plants. Further investigation of in vivo pmf partitioning to mediate the !! component as well as the regulation of photosynthesis by !! will likely identify key factor s that allow organisms to respond to the total pmf generated, as well as factors that allow organisms limit the impact of transient !! fluctuations experienced under natural conditions. 13 Chapter 2 Limitations to photosynth esis by proton motive force -induced photosystem II photodamage 1 Geoffry A. Davis, Atsuko Kanazawa, Mark Aurel Schottler, Kaori Kohzuma, John E. Froehlich, A William Rutherford, Mio Satoh -Cruz, Deepika Minhas, Stefanie Tietz, Amit Dhingra, David M. Kramer 1 1 This chapter was published as: Davis, GA, Kanazawa, A, Schottler, MA, Kohzuma, K, Froehlich, JE, Rutherford, AW , Satoh -Cruz, M, Minhas, D. Tietz, S, Dhingra, A, Kra mer, DM. 2016 Limitations to photosynthesis by proton motive force -induced photosystem II photodamage. eLife, 5. doi: 10.7554/eLife.16921 14 2.1 Abstract The thylakoid proton motive force ( pmf ) generated during photosynthesis is the essential driving force for ATP production; it is also a central regulator of light capture and electron transfer. We investigated the effects of elevated pmf on photosynthesis in a library of Arabidopsis thaliana mutants with altered rates of thylakoid lumen proton efflux, leading to a range of steady -state pmf extents. We observed the expected pmf -dependent alterations in photosynthetic regulation , but also strong effects on the rate of photosystem II (PSII) photodamage. Detailed analyses indicate this effect is related to an elevated electric field ( !!) component of the pmf , rather than lumen acidification, which in vivo increased PSII charge reco mbination rates, producing singlet oxygen and subsequent photodamage. The effects are seen even in wild type plants, especially under fluctuating illumination, suggesting that !!-induced photodamage represents a previously unrecognized limiting factor for plant productivity under dynamic environmental conditions seen in the field. 15 2.2 Introduction The thylakoid proton motive force ( pmf ), the transmembrane electrochemical gradient of protons generated during the light reactions of photosynthesis, is a fundamental entity of bioenergetics, coupling light -driven electron transfer reactions to the phosphorylation of ADP via the ATP synthase (59, 60). In oxygenic photosynthesis, light energy is captured by pigments in light -harvesting complexes and transf erred to a subset of chlorophylls in photosystem I (PSI) and photosystem II (PSII), where it drives the extraction of electrons from water and their transfer through redox cofactors to ultimately reduce NADP +. The vectorial transfer of electrons across the membrane is tightly coupled with the generation of the pmf , composed of both electric field (!!) and pH ( ! pH) gradients. In addition to its role in energy conservation, the pmf is also critical for feedback regulation of photosynthesis (61). Acidification of the thylakoid lumen activates the photoprotective energy -dependent exciton quenching (q E) process, which dissipates excess absorbed light energy in the photosynthetic antenna complexes by the activation of violaxanthin deepoxidase (62) and protonation of PsbS (7). Lumen acidification also regulates the oxidation of plastoquinol by the cytochrome b6f complex, slowing electron transfer from PSII and preventing the accumulation of electrons on PSI, which can otherwise lead to photodamage (11, 12). In vitro work has shown that excessive lumen acidification can inactivate PSII (63), decrease the stability of plastocyanin (64) and severely restrict electron flow through the cytochrome b6f complex (13). Taken together, the in vivo and in vitro evidence of the susceptibility of photosynthetic components to acidification has led to the proposal that the extent of pmf and its partitioning into !! and ! pH components is regulated to maintain the lumen pH above about 5.8, where it can regulate photopr otection. However, under environmental 16 stresses the regulation of photosynthesis may become overwhelmed, leading to PSII damage, or photoinhibition, from lumen over -acidification, although this has not been shown to occur in vivo (13). PSII photoinhibition can be a major contributor to loss of photosynthetic productivity (65), particularly under rapid fluctuations in environmental conditions experienced in the field (66). However, the mechanisms and regulation of photoinhibition remain highly debated (67, 68). Though several mechanisms have been proposed for the photodamage process, it is not known which of these operate in vivo under diverse environmental conditions. In addition, there are differing views on whether the extent of photoinhibition is governed by the rate of photodamage to PSII or by regulation of PSII repair (69). Answering these questions is essential to understanding how plants respond to rapidly changing conditions and thus of critical importance to improving plant productivity. The extent of the pmf can be modulated by altering the light -driven influx of protons into the lumen, i.e. by changing the rates of linear electron flow (LEF) or cyclic electron flow, or the efflux of protons through the ATP synthase (reviewed in 13). This latter mod e of regulation is important for co -regulation of the light reactions with downstream metabolic processes. For example, under CO 2 limitations (19) as well as during stromal P i limited conditions (18) the chloroplast ATP synthase activity is strongly down -regulated, decreasing the proton conductivity (gH+) of the thylakoid, leading to buildup of pmf and activation of q E. Similar decreases in gH+ and increases in pmf are seen when ATP synthase protein content is decreased (70) or in mutants with an altered AT P synthase "-subunit (71). 17 2.3 Results We took advantage of the effects of the ATP synthase on the photosynthetic proton circuit to probe how the pmf influences photoinhibition in vivo . We constructed a series of Arabidopsis thaliana mutants, which we termed minira (mini mum recapitulation of ATPC2), in which we complemented a "1-subunit (ATPC1) T -DNA knockout line (72) with "-coding sequences containing site -directed mutations to specifically incorporate amino acid changes around the redox regulatory cy steines present in ATPC2 into ATPC1 (Figure 2.1A) (73). Each minira line is designated numerically based on amino acid position and independent transformation events; for example minira 4Ð1, minira 4Ð2, and minira 4Ð3 represent three independent transforma tion events of the same I201V mutation. The minira library was originally developed as part of an on -going 'domain swapping' approach to assess functional differences in the two ATPC paralogs in Arabidopsis, ATPC1 and ATPC2, but the minira mutants also sho ws a range of ATP synthase activities useful to the present work. As described below, we also confirmed key aspects of the work using previously characterized ATPC1 tobacco antisense lines (70). However, variations in ATP synthase suppression in the antise nse lines between leaves and plant generations limited their experimental utility. The current mutagenesis approach was preferable as it allowed repeatable analyses of multiple photosynthetic parameters in stable, identical genetic backgrounds within each mutant line. 18 Figure 2.1 : "-subunit mutations alter photosynthetic proton efflux. Sequence alignment of Arabidopsis ATPC1 and ATPC2 regulatory region ( A). Amino acid differences incorporated into ATPC1 to generate minira are indicated by symbols ( & ). Amino acid numbers are based on standard spinach positions, which minira numeric designations are based upon position within the amino acid primary sequence. Regions where multiple changes were incorporated from ATPC2 are outlined in brackets. Bold: mutations resulting in successful transgenic plants with stable phenotypes utilized for experiments. Accumulation of chloroplast ATP synthase complexes was verified across resulting transformant lines ( B). Total leaf protein was probed for chloroplast ATP synthase %-subunit compared to a titration of wild type (Ws -2) accumulation. Gels run with identical samples stained with Coomassie Brilliant Blue are shown below to ensure equal loading to the 100% wild type samples. The conductivity o f the ATP synthase for protons ( gH+, C) and the light -driven pmf (ECS t, D), calculated from the decay of the electrochromic shift at 100 µmol photons m '2s'1 actinic light (mean ± s.d, n = 3). Statistically significant differences (*p<0.05) from wild type were determined using a t -test. ECS units were defined as the deconvoluted ! A520 µg chlorophyll '1 cm2. 19 We assessed the effects of minira modifications on gH+ and the extent of the light -driven pmf in vivo based on the decay kinetics of the electrochromic s hift (ECS), which reports changes in the thylakoid electric field (74). The minira lines displayed a range of ATP synthase activities (Figure 2.1C) , from about 30 Ð120% that of wild type (Wassilewskija -2, Ws -2), resulting in similar variations in light -driv en pmf (Figure 2.1D) . Multiple independent transformations were utilized for the same minira mutation, as the gH+ changes likely reflect both intrinsic ATP synthase activity changes due to the mutations, as well as changes due to protein expression level o r stability of the mutated subunit within the complex (Figure 2.1B) . While some minira mutants display an increase in gH+ relative to the wild type and will acidify the lumen at a slower rate, others have a large decrease in total ATP synthase content. For the purpose of understanding how a high pmf impacts photosynthesis we have primarily focused on those mutants that modified gH+ while maintaining an ATP synthase content similar to wild type levels. The minira library was then screened for photosynthetic phenotypes using whole plant chlorophyll fluorescence imaging (75) over a consecutive three -day photoperiod (Figure 2.2A) . Photosynthetic parameters were calculated from these images (videos are shown in supplemental files 1 -9) and shown as kinetic traces (Supplemental Figures 2.1 -2.16) or as log -fold changes compared to wild type ( Figure 2.2B -D). Under 'standard' laboratory growth chamber lighting on day one, most minira lines showed relatively small differences from wild type in LEF ( Figure 2.2B), qE (Figure 2.2C) , and photoinhibitory quenching (q I) (Figure 2.2D) , with the exceptions of minira 3Ð1 and 11 Ð1, which also showed the most severe decreases in gH+ (Figure 2.1C) . Stronger photosynthetic phenotypes appeared on days two and three, 20 implying that the decreased gH+ and pmf effects were enhanced by intense or fluctuating illumination. In general, LEF decreased with decreasing gH+ (Figure 2.2B , rows are ordered by increasing gH+) while q E increased ( Figure 2.2C) . The increases in q E were especially pr onounced at higher light intensities on days two and three. These effects can be explained by slowing of proton efflux through the ATP synthase in the mutants that results in increased pmf for a given LEF. This is reflected in the higher lumen pH -sensitive qE response for a given LEF ( Figure 2.2E) , which is strikingly similar to that attributed to ATP synthase regulation in wild type plants during limitations in carbon fixation (19) or decre ases in ATP synthase content (70). The qE sensitivities for the mutants remained similar throughout the experiments, i.e. the data for each mutant followed similar curves, implying that the ATP synthase activities were relatively constant within a particul ar line, consistent with a lack of light -dependent modulation of gH+ that has previously been observed (76). The extents of q E did not exceed about 3.5 units in any of the lines, suggesting that either the lumen pH was restricted to a moderate acidity or t hat the capacity of the q E response was saturated as light intensities increased. 21 Figure 2.2 : Dynamic light conditions enhance pmf dependent phenotypes. Whole plant fluorescent images were captured over three days under the illumination conditions displayed in Panel A and listed in Tables 2.5 -2.7. Plants were illuminated over the 16 -hr photoperiod, shown as yellow filled areas representing the light intensity when present in A, under either a constant light intensit y (day one), a sinusoidal photoperiod (day two), or a sinusoidal photoperiod interrupted by fluctuations in light intensity (day three). Square symbols in Panel A indicate each light intensity change. Steady -state fluorescence parameters were captured for each plant at the end of each light condition. Panels B, C and D represent the responses of LEF, q E and q I respectively (n ( 3). Data are shown as log 2-fold changes compared to the wild type. Kinetic data including wild type are shown in Supplemental Figur es 2.1 -2.16. The rows were sorted in order of ascending gH+ values measured as in Figure 2.1C . Panel E plots the dependence of the mean (n ( 3) of q E against the linear electron flow (LEF) for each time point measured for the day. For visualization purpose s the error bars have been omitted from E. 22 We also observed a strong correlation between increased q E and q I, (for quantitative comparisons, see Supplemental Figure 2.2) , implying that decreases in ATP synthase activity in the mutants led to not only highe r photoprotection but also higher rates of PSII photoinhibition. This result appears counterintuitive, in that we would expect the photoprotective q E response to prevent photoinhibition. Particularly striking was the relative loss in q E near the peak light intensities on days two and three in the most strongly affected minira lines ( Figure 2.2C) . We attribute this effect to strong accumulation of PSII photoinhibition ( Figure 2.2D) leading to the loss of photosynthetic capacity (see Figure 2.2B) that limited acidification of the thylakoid lumen. The observed increases in photoinhibition at high pmf could be caused by several mechanisms. It has been proposed that photoinhibition is primarily controlled by modulating the rate of PSII repair, i.e. the rate of da mage is dependent solely on light intensity but repair being inhibited by stress -induced ROS production (77). However, blocking PSII repair with the chloroplast translation inhibitor lincomycin (78) revealed that, when compared to wild type or minira lines with wild type like gH+ (e.g. minira 6Ð1), minira lines with low gH+ and high pmf (e.g. minira 3Ð1, Figure 2.1C , Figure 2.3) had higher rates of photodamage as reflected in both decreased maximal PSII quantum efficiency and loss of t he capacity to perform charge separation in PSII (79-81) and significantly decreased levels of D1 protein ( Figure 2.3C -E). Thus, decreasing ATP synthase activity led to increased PSII photodamage rather than decreased rates of repair, in contradiction with strict control of pho toinhibition by repair (69). 23 Figure 2.3 : Elevated pmf leads to PSII photodamage. Detached leaves were infiltrated with either water ( A, C) or a 3 mM solution of lincomycin ( B, D) and treated with 1000 µmol photons m '2s'1 red light for the times indicated . Following dark adaptation, F V/FM values were obtained ( A, B) for treated leaves of wild type (square, initial FV/FM 0.75 ± 0.006 and 0.76 ± 0.012 for water and lincomycin, respectively), minira 3Ð1 (triangle, initial F V/FM 0.60 ± 0.07 and 0.54 ± 0.10 for water and lincomycin, respectively), and minira 6Ð1 (hexagon, initial F V/FM 0.72 ± 0.004 and 0.73 ± 0.01 for water and lincomycin, respectively) (mean ± s.d., n ( 3). Data were normalized to the initial dark -adapted F V/FM values to remove intrinsic differ ences between the three lines. In panel B, dashed lines represent the best fit curves for a single exponential decay. The ability of photoinhibited leaves to perform PSII charge separation was determined in Ws -2 and minira 3-1 by measuring the ECS absorban ce changes following two consecutive single -turnover saturating flashes in the presence of DCMU ( C, D). Leaves infiltrated with water ( C, initial amplitudes of 8.88 ) 10'4 ± 9.0 ) 10'5 and 9.59 ) 10'5 ± 3.6 ) 10'5 for wild type and minira 3Ð1, respectively ) or 3 mM lincomycin (D, initial amplitudes of 8.59 ) 10'4 ± 1.2 ) 10'4 and 1.63 ) 10'4 ± 3.1 ) 10'5 for 24 Figure 2.3 ( continued ): wild type and minira 3-1, respectively) were infiltrated with DCMU following the indicated light treatment time and dark adapta tion. PSII activity was determined by subtracting the ECS amplitude induced by the second flash from the ECS amplitude induced by the first flash (mean ± s.d., n ( 4). Loss of the PSII reaction center D1 protein over the time course of illumination in linc omycin treated wild type and minira 3-1 leaves ( E). Leaves treated as in panel B were analyzed by western blot (n = 4) using an $-PsbA antibody and the 32 kDa band was quantified. Band intensities were normalized to the time zero point for each genotype within a single blot to control for differences in development intensities. Statistically significant differences (*p<0.05) from wild type were determined using a t -test. Photodamage can be induced in vitro by excitation of PSII centers with previously redu ced primary quinone acceptor (Q A) leading to the formation of the doubly -reduced QAH2 state (67). This situation might be expected if a high pmf slowed electron transfer through the b6f complex, resulting in the accumulation of electrons on PSII acceptors. When data at a range of light intensities are compared, the relationship between the Q A redox state (q L) (82) and qI is statistically significant (ANOVA, p=2 ) 10-16) (Figure 2.4) . However, both the light intensity (p=3 ) 10-5) and q E (p=7 ) 10-3) are sig nificant interacting factors. At any one light intensity, q L was relatively stable, whereas q I was strongly dependent upon the mutant background and underlining pmf changes, indicating that while Q A reduction may be a contributing factor, it cannot by itse lf explain the observed extents of photoinhibition in the minira lines. On the other hand, this dependence is also consistent with an alternative model, proposed below, that involves effects on PSII recombination rates. 25 Figure 2.4 : The dependence of phot oinhibition on the redox state of Q A. The redox state of the primary electron acceptor Q A was assayed using the q L fluorescence parameter concurrently with photoinhibitory quenching q I at 100 (solid symbols), 300 (half filled symbols), and 500 µmol photons m'2s'1(open symbols, mean ± s.d., n = 3). Plants were exposed to at least 10 min of actinic illumination prior to q L measurement, and q I measured after 10 min of dark relaxation. While the extent of q I varies between plants, the relative redox state of QA remains similar between all plants within each actinic light intensity. Changes in chlorophyll content have also been correlated with increases in PSII photoinhibition (83), likely due to less light being absorbed at the leaf surface and the subsequent increased light penetration into the leaf reaching more PSII centers. While the leaf chlorophyll content was altered in the minira mutants from wild type levels ( Table 2.8) , the leaf chlorophyll content does not fall below where (83) observed correlations between a lack of chlorophyll content and photoinhibition. We next hypothesized that the most probable explanation for the increased photoinhibition is direct sensitization of PSII to photodamage by pmf . In the 'acid -damage' model (13), it was proposed that excessive lumen acidification at high ! pH (i.e. low lumen pH) could sensitize PSII centers to photodamage. To test this possibility, we compared the rates of 26 photoinhibition with the extents of the ! pH and !! components of the pmf by measuring the relaxation kinetics of the ECS signal (47). Surprisingly, increased extents of photoinhibition in low gH+ lines were not correlated with ! pH, but were with !! (Figure 2.5) , contrary to what was expected with the acid -damage model. Consiste nt with this result, the rates of P 700+ reduction, which reflect the lumen pH -sensitive turnover of the cytochrome b6f complex remained in a range consistent with a lumen pH above or near the p Ka for b6f down -regulation, i.e. above about 6.0 (Supplemental Figure 2.17) . Figure 2.5 : Photoinhibition is strongly correlated with !! but not !pH in minira lines. Photoinhibition, estimated by the q I fluorescence parameter, is plotted against either the ! pH or !! components of pmf , estimated by the ECS ss (A) and E CSinv (B) parameters, as described in Materials and methods. Measurements shown were taken during exposure to 500 µmol photons m'2s'1 actinic light (mean ± s.d., n = 3). Two -way analysis of variance (ANOVA) of all combined data, 15 minira lines and wild type, showed a stronger correlation between q I and !! (F = 9.5, p=0.003) than ! pH ( F = 4.05, p=0.05). This correlation is also seen with the expected pH-dependent alterations of P 700+ reduction for the observed partitioning differences from wild 27 Figure 2.5 (continued ): type ( Supplemental Figure 2.17) and an increase in the fraction of total pmf stored as !! at the expense of ! pH over multiple light intensities ( Supplemental Figure 2.18). Increased storage of pmf as !! is also observed in tobacco ATP synthas e knock -down plants ( Supplemental Figure 2.19) . ECS units were defined as the deconvoluted ! A520 µg chlorophyll '1 cm2. The influence of !! on the rate of PSII recombination was estimated based on the change in the equilibrium constant for the sharing of e lectrons between pheophytin and QA (described in Materials and methods) ( C). The influence of !! on the calculated recombination rate taking into account the fraction of reduced Q A using the equations described in Materials and methods and described in the main text. Data were obtained at 100 (solid symbols), 300 (half filled symbols), and 500 µmol photons m '2s'1 (open symbols) (mean ± s.d., n = 3). These results are consistent with an increase in the partitioning of pmf into !! as the total pmf increased (Supplemental Figure 2.18) , as have been observed previously (59), and are likely caused by alterations in ion movements across both the thylakoid and chloroplast envelope membranes (28, 29, 59). Our results suggest that pmf partitioning acts to maintain a permissible lumen pH during large pmf increases. We observed similar results throughout the range of minira mutants with low gH+ as well as with tobacco "-subunit antisense plants (70) with reduced ATP synthase complexes, finding that a large increase in total pmf was accompanied by increased partitioning of pmf into !! (Supplemental Figure 2.19) , and increased rates of photodamage in the presence and absence of lincomycin ( Supplemental Figure 2.19) . These results suggest that low gH+ or high pmf -related PSII damage is a more general phenomenon, which is related to excess !! and likely to be independent of such factors as changes in the protein or supercomplex content (see also discussion in (70). As described in Figure 2.8 , we hypothesize that high !! accelerates photodamage by favoring recombination reactions within PSII (55, 84-86) that lead to the formation of chlorophyll triplet states that in turn generate 1O2 (87). 28 To test this model, we studied the relationship between elevated !! and PSII charge rec ombination in isolated spinach thylakoids ( Figure 2.6) , which unlike Arabidopsis can be isolated as highly intact chloroplasts and tightly coupled thylakoids. Consistent with our model, we found that elevated !!, produced by artificial decyl -ubiquinol medi ated cyclic electron flow through PSI, increased the rate of charge recombination from the S 2QA- state compared to samples treated with gramicidin to dissipate !! (Figure 2.6A) . This recombination reaction can also occur in vivo during normal turnover when the S 2QA- state is formed given that the equilibrium constant for sharing of electrons between Q A and Q B is small (88). The extent of !! generated in these experiments was similar to that observed in minira leaves under photoinhibitory conditions ( Figure 2.6B), suggesting similar increases in recombination rates should occur in vivo . Figure 2.6 : In vitro manipulation of the !! alters PSII S 2QA- recombination rates. Isolated spinach thylakoids in the presence of 5 µM spinach ferredoxin and 10 µm sodium ascorbate were treated with 3 Ð3,4-dichlorophenyl 1,1 -dimethylurea (DCMU) to block PSII forward electron transfer, and a trans -thylakoid pmf generated utilizing decyl -ubiquinol mediated PSI cyclic electron transfer ( A). Recombination from the S 2QA' state was p robed by observing the decrease in the high fluorescence state associated with Q A' following a short (100 ms) actinic flash to dark -adapted thylakoids (black line). Depletion of the pmf , which under these conditions is stored almost exclusively as !!, in t he presence of 25 µM gramicidin (red line) resulted in an approximate 5 -fold increase in the initial rate of decay when !! was largest, and an overall 2 -fold increase in the lifetime of the high fluorescence state. The extent of !! generated by the 100 29 Figure 2.6 (continued ): ms light ( B), estimated by the ECS signal measured at 520 nm and normalized to chlorophyll content. Thylakoids were assayed in the absence of inhibitors (green line), in the presence of DCMU (blue line), and in the presence of both DCMU and 50 µM decyl -ubiquinol (black line) to generate pmf through PSI turnover, corresponding to the condition used in panel A. We next tested the predicted connection between elevated !! and singlet oxygen ( 1O2) generation ( Figure 2.7D) . In wild type l eaves, moderate illumination (30 min of 300 µmol photons m -2s-1) resulted in no detectable light dependent changes in 1O2 (Figure 2.7D) using Singlet Oxygen Sensor Green (SOSG) dye fluorescence (89-91). In contrast, the low gH+ minira 3Ð1 line showed a str ong induction of SOSG fluorescence within the first 10 min of illumination, which saturated by about 30 min. Infiltration of leaves with valinomycin, a potassium ionophore that decreases the !! component of pmf (85), partially inhibited the rise in in SOSG fluorescence ( Supplemental Figure 2.20) . While care must be taken making quantitative estimates of 1O2 from SOSG fluorescence (92), within the limits of these experiments the !!-dependence of the SOSG fluorescence increases strongly support a role for !!-induced 1O2 production from photosynthesis. 30 Figure 2.7 : Induction of !! and 1O2 production under fluctuating light in wild type plants. (A) Illumination conditions and measurement points used in the experiments. Fluctuating light conditions (replicating Figure 2.2 day three) are shown as connected points, with open squares representing measurements obtained 10 s after the light transition and closed squares the end of steady -state illumination. Constant illumination of 300 µmol m -2s-1 is represented as a dotted line. ( B) Representative traces of the light -fluctuation induced ECS signals resulting in transient ECS 'spikes' are shown for the first fluctuation (dark to 39 µmol m '2s'1, blue) and the fluctuation from 167 to 333 µmol m '2s'1, red). A full set of ECS kinetic 'spikes' following increased light fluctuations can be found in Supplemental Figure 2.21 . The extents of light -induced !! in wild type (green line and shaded box, indicating mean ± s.d) and minira 3-1 (red lines and box, indicating mean ± s.d) at 300 µmol m '2s'1 are shown for comparison. ( C) The extents of light -induced !!, estimated using the ECS ss parameter over the time -course of the fluctuating light experiment, compared to those obtained under continuous illumination in wild type and minira 3-1 (green and red lines and boxes, respectively, as in Panel B). Open and closed squares correspond to the ECS ss measurements taken at the timing designated in panel A. (D) Time -course of SOSG fluorescence changes for wild type (squares) and minira 3Ð1 (triangles) during exposure to constant 300 µmol m '2s'1 (dotted lines) and wild type leaves under the first hour of fluctuating light (solid line). A decrease in SOSG fluorescence occurs when !! is collapsed with the addition of the ionophore valinomycin ( Supplemental Figure 2.20) . All data in A, B, and C represent mean (n ( 3) ± s.d. ECS units were defined as the deconvoluted ! A520 µg chlorophyll '1 cm2. 31 Singlet oxygen is produced by the interaction of O 2 with triplet excited states of pigments, most likely from the 3P680 chlorophyll within the PSII reaction centers generated by recombination reactions (93, 94), as further discussed below. It is unlikely that such triplets could be generated in the bulk light harvesting pigments because high NPQ in the mutant s will decrease the lifetime of antenna excited states, and chlorophyll triplets generated in light harvesting complexes are efficiently quenched by carotenoids (95). We thus propose that elevated !! induces 1O2 production by accelerating PSII recombinatio n in low gH+ mutants when pmf is large. Keren et al. (96) suggested that recombination -induced triplet formation could explain the photoinhibitory effects of very low light, when PSII charge recombination is preferred over the forward electron transfer rea ctions. Our work implies that this type of phenomenon is greatly accelerated by high !!, potentially making it relevant to photosynthesis under growth light conditions. The effects of high !! would be expected to alter the rates of PSII recombination throu gh P+Pheo - (where P + is the oxidized primary chlorophyll donor and Pheo the D1 subunit pheophytin) in an increasing dependence upon the fraction of Q A-, consistent with the observed correlation between q I and q L as the light intensity increased ( Figure 2.4 ). To test this relationship, we estimated the recombination rates from S 2QA- through the P +Pheo - pathway, considering Q A redox state (estimated by 1 -qL) and the expected impact of !! on the equilibrium constant for sharing electrons between Pheo and Q A (based on ECS ss and the position of Q A in the structure relative to the membrane dielectric). The basis of this estimate is described in more detail in Materials and methods. As shown in Figure 2.5C , we see a positive correlation between qI and estimated recombination through P +Pheo - over both mutant variants and light intensities, indicating that the combined effects of !! and Q A redox state can explain a large fraction of the 32 observed extents of photoinhibition. While it is likely that multiple mechanism s of photoinhibition exist, which may also explain some of the q I variation, overall the greatest impact upon photoinhibition under these conditions can be explained by !!-mediated changes in PSII electron recombination. The obvious questions are: does !!-induced photoinhibition occur in wild type plants and if so under what conditions? Photosynthesis is known to be particularly sensitive to rapid fluctuations in light intensity (66), at least some of this sensitivity is associated with photoinhibition of P SI, especially in cyanobacteria (97). However, such fluctuations should also result in large transient changes in !!, as the thylakoid membrane has a low electrical capacitance and low permeability to counter -ions, while the lumen and stroma have high prot on buffering capacity (2). The slow onset of q E and other down -regulatory processes in photosynthesis should exacerbate these effects and allow for large fluxes of electrons when light levels are rapidly increased, resulting in large, transient !! 'pulses' . We therefore hypothesized that !!-induced photodamage may contribute to the increased photodamage seen under fluctuating light. To test this possibility, we measured light -driven pmf (!! and ! pH) and 1O2 generation in wild type Arabidopsis under fluctuat ing light conditions ( Figure 2.7A , replicating the first 8 * hr of Figure 2.2A day three). The initial dark -to-light transition resulted in an immediate, transient 'spike' in !!, even though the light intensity was low (39 µmol photons m -2s-1, Figure 2.7B) . The spike was transient, and decreased to steady -state levels within tens of seconds, as shown in the blue trace in Figure 2.7B . Spikes in !! of similar amplitudes were also seen upon each increase in light at the onset of each fluctuation, though the reco very kinetics tended to be more rapid than those seen at the first dark -light transient. An example of these transients, taken at the transition between 167 and 333 µmol 33 photons m -2*s-1 is shown in ( Figure 2.7B) . A more complete set of transient kinetics, over the entire course of the experiment is presented in Supplemental Figure 2.21. The amplitudes of these !! transients were similar to or larger than those seen in the minira 3Ð1 line under constant 300 µmol photons m -2*s-1 light, which also induced 1O2 generation (see red horizontal bars in Figure 2.7B) . These spikes reflect increases in !! above that already produced by steady -state photosynthesis, so the true extent of !! is likely considerably higher, and based on estimates of the calibration of the ECS signal likely range between 150 Ð260 mV (see Supplemental Figure 2.21). By contrast, wild type plants under constant 300 µmol photons m -2*s-1 produced much lower !! extents ( Figure 2.7B green bars) and had no detectible 1O2 generation ( Figure 2.7D) . The se results suggest that even low amplitude light fluctuations are capable of inducing !! large enough to produce 1O2 and PSII photodamage. Supporting this interpretation, wild type leaves under these fluctuating light conditions produced substantial amount s of 1O2 during the first hour of fluctuating conditions compared to higher intensity, but constant illumination ( Figure 2.7D) . The above results lead us to conclude that fluctuating light likely induces strong effects through !!-induced recombination reac tions in PSII (see below for considerations of potential contributions to PSI). In addition to direct damage to the enzymes of photosynthesis, 1O2 can also activate plant light stress -related gene expression and programmed cell death (98), suggesting a pos sible physiological linkage between pmf -enhanced recombination and plant regulatory pathways that may result in long -term acclimation to fluctuating light. At a mechanistic level, we propose that imposing a large !! across the PSII complex will decrease th e standard free energy gap between the vectorial electron transfer steps and thus the back -reaction will be accelerated in competition with forward (energy -storing) reactions (37, 55, 34 87) (Figure 2.8) . It is known that decreasing the energy gap between Phe o and Q A favors recombination from P +QA- via Pheo - (55, 87) rather than directly from Q A- to P + (56, 87, 99). The observed increased recombination rates in vitro (Figure 2.6) when !! is present, combined with 1O2 production under high !! conditions in vivo suggest that the fraction of electrons recombining to P + through a Pheo - intermediate is greatly increased, as the recombination pathway via the Pheo - has a high yield for formation of the triplet state of P ( 3P) that in turn can interact with O 2 to produce 1O2 (56, 67, 87). 35 Figure 2.8 : Schemes for the trans -thylakoid !!-induced acceleration of recombination reactions in PSII and subsequent production of 1O2. (A) The relative positions of PSII electron transfer cofactors with respect to the electr ic field (double -headed arrow) imposed across the thylakoid membrane (dotted lines). The red and blue arrows indicate the !!-induced changes in the equilibrium constant for the sharing of electrons (') between Q A and Pheo, and electron holes (+) among P + and the oxygen evolving complex. Excitation of PSII by light leads to formation of excited chlorophyll states (P*), the excitation is shared over the 4 chlorophylls and the 2 pheophytins. Charge separation occurs between more than one pair of pigments, so a t short times the situation is not well defined, but ChlD1+Pheo D1' appears to be the dominant radical pair. Secondary electron transfer events occur forming P D1+Pheo D1', the second radical pair, which is present in nearly all centers. This radical pair is stabilized by electron transfer from Pheo ' to Q A forming P D1+QA'. This radical pair is further stabilized by electron transfer from D1Tyr161 (TyrZ) forming a neutral tyrosyl radical, 36 Figure 2.8 (continued ): which oxidizes the Mn cluster of the oxygen evolving complex to form the state S n+1 QA'. Finally, Q A' reduces Q B to form S n+1 QB-. Upon a second PSII turnover the double reduced and protonated Q B plastohydroquinone becomes protonated and is exchanged with an oxidized plastoquinone from the membrane pool (black arrows). The illus tration was based on crystal structure 3WU2 ( Umena et al., 2011 ). (B) The charge separation states described above are unstable and recombination competes with the energy -storing reactions . When P D1+ is present, recombination reactions can occur by several pathways as indicated by the dashed lines: (1) direct electron transfer from Q A' to P + (R1); (2) by the back reaction to form the P+Pheo ' state, which can then recombine directly (R2) or, (3) when the P +Pheo ' radical pair is present as a triplet state, 3[P+Pheo '], the dominant state when formed by the back -reaction, 3[P+Pheo '] charge recombination forms 3P (R3), a long lived chlorophyll triplet that can easily interact with O 2 to form 1O2; (4) complete reversal of electron transfer can also occur, repopulating P* (R4), which can return to the ground state by emitting fluorescence (luminescence) or heat. Route 3, the triplet generating pathway, is the dominant recombination route in fully fu nctional PSII. For simplicity the 3[P+Phe -] is not distinguised from the singlet form in this scheme. A !! across the membrane should destabilize P +QA' relative to the other states (see dotted blue lines), affecting the rates of reactions indicated in the green versus red. A !! across the membrane should also destabilize P +Pheo ', but because of the smaller dielectric span across the membrane, to a lesser extent than P +QA'. Thus, the buildup of !! should shift the equilibrium constant for sharing electrons b etween Q A' and Pheo, favoring the formation of Pheo ' and thus increasing the rate of recombination through R3 (as well as the R2 and R4), resulting in increased production of 3P and 1O2. Destabilization of P +QA' will also increase the driving force for P +QA' recombination via R1, however this recombination is already driven by 1.4eV and it is thus likely to be in the Marcus inverted region. Thus increasing the driving force will slow recombination by this route. 2.4 Discussion This proposed mechanism of !!-mediated photoinhibition has broad implications for the energy limitations of photosynthesis. Although the two components of pmf are energetically equivalent for driving ATP synthesis (3), they have distinct effects on the regulation of photosynthesis (13, 100, 101). It has thus been proposed that the partitioning of pmf into !! and ! pH is regulated to maintain a balance between efficient energy storage and regulation of light capture (61). We demonstrate here another important constraint on this balance: the avoidance of photodamage caused by recombination reactions in PSII, and this may explain the need for complex ion balancing systems in chloroplasts (2, 28, 29). 37 The effect of !! on recombination and 1O2 production may, at least in part, explain the severe effects of fluctuating light on photosynthesis, and thus could constitute a significant limitation to photosynthetic productivity. In the absence of a large !!, the energy gap between the P +Pheo - and P +QA- appears to be sufficient to keep detrimental recombination to a manageable level when the usual regulatory mechanisms are functional (56). During photosynthesis, and especially under fluctuating light, though, the energy gaps between the photo -generated r adical pairs vary dynamically under the influence of the pmf , so that high !! renders the charge -separated states in the photosystems considerably less stable. The !! is also expected to influence the trap depth (i.e. the energy level between P * and the fi rst radical pair(s), the most relevant probably being P +Pheo -) and this could potentially affect the quantum yield of charge separation and the yield of radiative recombination (luminescence). Transthylakoid electric fields also influence recombination rea ctions in PSI reaction centers (102). It is thought that P 700 triplet formation is minimized in PSI by the presence of the higher potential quinone in PsaA, making this side of the reaction center the safe charge recombination pathway (56). In light of the present findings, it is worth considering whether a transiently large !! could make this protective mechanism less efficient, though this question has yet to be addressed experimentally. Over evolutionary time scales, the !!-effect may have constrained ot her bioenergetics features of photosynthesis. It has long been known that oxygenic photosynthesis is limited, in most organisms, to wavelength ranges shorter than about 700 nm (the 'red limit'), resulting in the loss of a large fraction of the light energy hitting the plant (103). Gust et al. (104) proposed that the red limit may be the result of certain limitations imposed in part by key biochemical properties of life (e.g. the properties of energy storage molecules NAD(P)H and ATP, the use of 38 certain bioc hemical pathways, etc.) that evolved before the advent of photosynthesis. Milo (105) came to a different conclusion based on estimates of the theoretical wavelength dependence of energy conversion efficiency for plant photosynthesis, based on the Shockley and Queisser equation (106). The maximum efficiency was about 700 nm, similar to the red limit of oxygenic photosynthesis, suggesting that evolution has selected for photosynthetic energetics based on this fundamental limit, rather than any biological impe rative. However, the predicted wavelength -dependence of the energy efficiency is very broad, with only about a 15% decrease from the peak at wavelengths out to 800 nm, far beyond the red limit, even in organisms with red -shifted reaction centers. Marosvolg yi and van Gorkom (107) drew a similar conclusion but with additional restraints and suggested a narrower maximum more closely overlapping with the red absorption of chlorophyll a. Rutherford et al. (56) took a different view based on an analysis of the bi oenergetics of reaction centers. They noted that PSII was in a uniquely difficult situation in energy terms: (1) it does multi -electron chemistry (at both sides of the reaction center) and so cannot prevent back reactions by kinetic control, (2) it has a v ery energy demanding reaction to do: water oxidation and quinone reduction with a + E ~920 meV in functional conditions, and requires a significant over -potential not just for attaining a high quantum yield of photochemistry but also for achieving water oxi dation and quinol release, and (3) its chlorophyll cation chemistry is uniquely oxidizing and thus it cannot use carotenoids to protect itself from chlorophyll triplet formation at the heart of the reaction center. This situation means that unlike other ty pe-II reaction centers, PSII is unable to prevent electrons from the bound semiquinones getting back to the Pheo and then recombining with P + and forming 3P. 39 This lack of energy 'headroom' was seen not only as a major factor in PS IIÕs susceptibility to ph otodamage but also as a reason why oxygenic photosynthesis is pinned to chlorophyll a photochemistry at around 680 nm as the red limit. The existence of efficient oxygenic photosynthesis at longer wavelengths seemed to question that view (108-111). However , it was pointed out that these species seem to exist in very stable environments that have very little variation in light conditions (112). Under such a narrow range of illumination conditions it is not unreasonable that less energy 'headroom' is required (112). Clearly, these specific energy limitations of PSII will be exacerbated by spikes in the !! reported here. Indeed, it seems reasonable to suggest that the existing 'energy headroom' postulated in normal chlorophyll a-containing PSII, while too small to avoid photodamage altogether, exists quite specifically to mitigate the extra photodamage from back -reactions enhanced by spikes in !! due to variable light intensities. In this way, the extent of the variable light -induced !! may be considered to cont ribute to the position of the red limit of oxygenic photosynthesis. The need to prevent !!-induced recombination may also have guided the evolution of other photosynthetic components. Recent mechanistic models of the ATP synthase suggest that the ratio of protons passed through the ATP synthase per ATP synthesized depends on the number of subunits in the c-ring (41). The chloroplasts of green plants and algae thus far studied possess ATP synthase complexes with larger c-ring stoichiometries than their mitoc hondrial and bacterial homologues (45), imposing higher fluxes of protons to generate ATP and necessitating the engagement of additional bioenergetic processes, including cyclic electron flow, to make up the ATP deficit needed to sustain photosynthesis (76, 82). While this increased H + demand is often viewed as a bioenergetic limitation to photosynthetic electron and proton transfer, a high 40 H+/ATP ratio decreases the pmf needed to maintain a given ATP free energy state ( ! GATP ), thus allowing photosynthesis to operate at a decreased steady -state pmf . In this context, the deleterious electron recombination effects of a high pmf may have favored the evolution of ATP synthase complexes with high H +/ATP ratios in chloroplasts. 2.5 Materials and Methods 2.5.1 Plan t materials and growth conditions. Wild type Arabidopsis thaliana (ecotype Wassilewskija -2) and ATP synthase "-subunit mutants were germinated on Murashige and Skoog medium supplemented with 2% (w/v) sucrose, and 10 mg L -1 sulfadiazine for selection of tra nsgenic minira lines (113). Following germination plants were grown on soil under a 16 * hr photoperiod at 100 µmol photons m -2*s-1 at 22¡C for three weeks. Nicotiana tabacum wild type (cv Samsun NN) and ATPC1 antisense lines were germinated and grown as in (70) under a 16 * hr photoperiod at 300 µmol photons m -2*s-1. Measurements were performed at the onset of flowering on the youngest, fully expanded leaves. 2.5.2 Generation of chloroplast ATP synthase "-subunit minira (minimum recapitulation of ATPC2) mutant s. A T -DNA insertion mutant for ATPC1 ( dpa1) (72) was used to introduce the mutated constructs as a complemented allele as in (71). Site -directed mutations in the redox -regulatory domain of ATPC1 were designed to incorporate amino acid differences from the redox inactive ATPC2 into the redox regulated ATPC1 ( Figure 2.1A) . The Arabidopsis thaliana AtpC1 gene was excised from binary vector pSex001 as a SmaI/XbaI fragment and cloned into SmaI/XbaI digested pBluescript plasmid. The resulting plasmid was named p DA15. The mutations were introduced into ATPC1 using a combination of three approaches. 41 The first approach used an adaptor ligation strategy. Oligonucleotides were designed to introduce desired mutations. Adaptors representing the 5Õ and 3Õ strand of DNA t argeting specific mutations were obtained independently from Sigma Aldrich. A total of 5 * ml of each oligonucleotide pair (10 * mM) were mixed and denatured at 95¡C for 10 min in a boiling water bath. The oligonucleotides were allowed to reach room temperatur e over two hours in the water bath allowing for efficient annealing of complementary strands. The adaptors thus obtained were utilized for adaptor ligation. The target region from ATPC1 cDNA was removed using BglII/HpaI restriction enzymes. The larger line arized backbone of pDA15 was used for ligation with the adaptors (Table 2.1) . The resultant vector was double digested with SmaI and XbaI to excise the mutated ATPC1 gene, which was then ligated back into the SmaI/XbaI digested binary vector, pSex001. A se cond set of mutations was introduced by first digesting pDA15 with BglII followed by a partial digestion with TatI. The resulting plasmid backbone was used for ligation with the adaptors having the desired mutation ( Table 2.2) . As in the first adaptor liga tion -mediated mutagenesis approach, the resultant vector was double digested with SmaI and XbaI to excise the respective mutated atpC1 gene and ligated into SmaI/XbaI digested binary vector, pSex001. A second mutagenesis strategy used splicing by overlap e xtension (SOE) PCR. For each desired mutation, two sets of primers were designed to produce two overlapping fragments during amplification of atpC1 from pDA15 such that the mutation was generated in the region of overlap ( Table 2.3) . The two fragments were mixed together and the resulting DNA solution was used as a template for a subsequent PCR using primers that amplify the complete AtpC1 gene (DMP 45 and DMP 46). This led to the amplification of the entire atpC1 gene with the desired 42 mutation. The mutated atpC1 was digested with SmaI and XbaI and was sub -cloned into SmaI/XbaI digested pSex001. The third mutagenesis approach involved swapping of target domains (delete swaps) using synthetic gene fragments synthesized at GenScript USA ( Table 2.4) . The synthe tic gene was used to replace the ATPC1 redox regulatory domain in the native gene. To swap the domains, pDA15 was digested with BsrGI and XbaI to remove the native domain and ligated with the synthetic fragment derived from the synthetic gene construct aft er digestion with BsrGI and XbaI. To introduce a synthetic gene with a single nucleotide mutation, the synthetic gene construct for minira 3 and pDA15 were double digested with BsrGI and XbaI and the synthetic BsrGI/XbaI fragment with the mutation was liga ted into pDA15 where the original BsrGI/XbaI fragment had been removed The resultant intermediate plasmid was digested with SmaI/XbaI to obtain the gene with swapped C domain. This gene was then sub -cloned into SmaI/XbaI digested pSex001. All of the introd uced mutations were confirmed by Sanger sequencing of individual plasmids. Following successful mutagenesis, minira constructs were mobilized into the binary vector pSEX001 -VS under control of the Cauliflower mosaic virus 35S promoter. A single minira cons truct was transformed into heterozygous dpa1 plants via Agrobacterium tumefaciens -mediated transformation (114). The resulting transgenic plants were screened for the minira insertion via PCR using the forward primer 5Õ -GGTAATATCCGGAAACCTCC - 3Õ and the reverse primer 5Õ -GTACAAGAGCTCGACTTTCTCG - 3Õ followed by dpa1 screening using the forward primer 5Õ -CACATCATCTCATTGATGCTTGG - 3Õ and the reverse primer 5Õ -GTACAAGAGCTCGACTTTGTCG - 3Õ. Transgenic plants containing both minira and dpa1insertions were self -pollinated until plants were homozygous for 43 both dpa1 (!atpc1 ) and the correct minira mutation, which was subsequently confirmed by sequencing. 2.5.3 Isolation of tightly coupled chloroplasts and intact thylakoids. Chloroplasts were extracted from market sp inach with modifications to the method described in Seigneurin -Berny et al. (115). All centrifugation steps were carried out at 4¡C and exposure to light was kept to a minimum. Briefly, approximately 20 * g of spinach leaves were homogenized in a blender for 10* s with ice cold homogenization buffer of 50 * mM HEPES (pH 7.6), 330* mM sorbitol, 5 * mM MgCl 2, 2* mM EDTA and supplemented with 0.1% BSA for grinding. The homogenate was filtered through three layers of wetted Miracloth and one layer of wetted muslin follo wed by centrifugation at 4000 x g for 10 * min. The pellet was resuspended in homogenization buffer and layered on top of a single step 80% Ð40% Percoll gradient. Intact chloroplasts were recovered after centrifugation for 20 * min at 3000 x g in a swinging buc ket rotor. The intact chloroplasts were diluted approximately 4 -fold with homogenization buffer and centrifuged for 5 * min at 4000 x g. The chloroplast pellet was resuspended in a minimal amount of homogenization buffer (<1 * mL) and chlorophyll quantified in 80% acetone (116). 2.5.4 Spectroscopic measurements. Near simultaneous chlorophyll fluorescence and electrochromic shift (ECS) measurements were performed on a custom made spectrophotometer (117). For in vivo spectroscopic measurements, following a 10 * min dark acclimation the maximal PSII quantum efficiency, linear electron flow (LEF), energy -dependent exciton quenching (q E), and photoinhibitory quenching (q I) were estimated using saturation pulse chlorophyll a fluorescence as described previously (118). The extent of the q I component of NPQ was determined following at least 10 * min dark relaxation to eliminate the residual effects of q E type quenching. Red actinic 44 illumination was used for all measurements to prevent incorrect assessment of chloroplast move ment as q I, as red light is ineffective in inducing chloroplast movements (119). A Stern -Volmer derivation of q E (qE(SV) ) was used to minimize the contribution of q I in the determination of q E (120). Estimates of the relative redox status of Q A (qL) were p erformed as described in (100) after at least 10 min of actinic illumination. The relative extents of steady state pmf (ECS t) and the conductivity of ATP synthase to protons ( gH+) were measured using the dark interval relaxation kinetics of absorbance changes associated with the electrochromic shift (ECS) fit to a first -order exponential decay (74). Partitioning of the pmf was determined from deconvolution of the absorbance change at three wavelengths (505, 520 and 535 nm) around 520 nm during the dark interval ECS changes and the ECS steady -state ( !!) and ECS inverse ( ! pH) were determined as in (47). Briefly, the total amplitude of the deconvoluted ECS signal following a rapid lig ht/dark transition was used to estimate the total light -induced pmf (ECS t). The steady -state !! component was determined from the extent to which the inverted ECS signal during the dark interval decreased from the steady -state baseline. The steady -state ! pH component was determined as the amplitude of the inverted ECS signal during the dark interval. The ECS measurements were corrected for pigment variations by normalizing to chlorophyll content determined from acetone extraction as above. For tobacco measu rements, the ECS measurements were normalized to the xenon -flash induced extent of the ! A520 nm ECS rise. P 700+ reduction kinetics were measured from the dark interval relaxation kinetics of the absorbance change at 810 nm after subtracting the 930 absorba nce change (121). In vitro chloroplast measurements were performed on a similar instrument described above modified to measure a cuvette held sample. Chloroplasts were osmotically shocked on ice in buffer containing 10 * mM HEPES (pH 7.8) and 10 * mM MgCl 2 to a final chlorophyll 45 concentration of 20 * µg ml -1 supplemented with 5 * µM spinach ferredoxin and 10 * µM ascorbate. Where noted, thylakoids were treated with 50 * µM decyl -ubiquinol to catalyze PSI cyclic electron transfer and generate a pmf , 50* µM 3 -(3,4-dichlor ophenyl) -1,1-dimethylurea (DCMU) to block PSII forward electron transfer, and 25 * µM gramicidin to decouple the pmf . Fluorescence measurements were performed as above, with variable fluorescence measured after a 20 * min dark adaptation after which the thylak oids were excited with a single 100 ms subsaturating actinic pulse. From a dark -adapted state, the application of a single turnover pulse will lead to rapid accumulation of !!, due to the high buffering capacity of the thylakoid lumen as well as the low capacitance of the thylakoid membrane, allowing gramicidin to decouple !!, as the !! primarily composes the pmf under these conditions (2). The F 0 measurements for all samples were taken from the first measured point to avoid any actinic effects due to the m easuring pulses themselves. 2.5.5 Estimation of recombination rate. The !!-induced enhancement of the rate of recombination from the S 2QA- state was estimated based on the change in the equilibrium constant for the sharing of electrons in the presence of !!. Other states will also recombine (e.g. the S 3QA- state) but we use S 2QA- as a proxy because we expect most of these states to respond to changes in Q A and !! in similar ways. The rate of recombination from S 2QA- was calculated as: !!!!!!!!!!!!!"!!!!"#$ !!!"!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!( 1 ) where [S 2QA-] is the concentration of PSII centers with reduced Q A, kr the intrinsic rate of recombination from S 2QA-, and ! Estab is the free energy for stabiliza tion of the charge separated state, expressed in eV. In the absence of a field and in the presence of DCMU where all Q A is reduced, vr is measured to be about 0.3 * s-1 (Figure 2.5) , but in the uninhibited complex under 46 steady state photosynthesis, this rate will be decreased proportionally by oxidation of Q A, while !Estab will be decreased by !! so that: !!!!!!!!!!!!!"!!!!"# !!!!"#$ !"!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!( 2 ) where 1 -qL is an estimation of the frac tion of Q A in the reduced form (100), and !!light -dark is the light -dark difference in electric field in mV. From Takizawa et al. (47) we obtained a factor for estimating !! from the extents of ECS, and correcting for the chlorophyll content as performed here, we obtain: !!!!!!!!!!!!!"!!"#!!!"!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!( 3 ) where ECS ss is the normalized ECS signal representing the light -dark difference in !! normalized to the chlorophyll content. 2.5.6 Chlorophyll fluorescence imaging. In vivo whole plant chlorophyll a fluorescence imaging was performed in an imaging chamber equipped with 50W Bridgelux White LEDs (BXRA -56C5300, Bridgelux Inc., Livermore, California) for white actinic illumination (75). Pre -illumination values for F 0 and FM were captured just prior to the beginning of the photoperiod and subsequent fluorescence parameters obtained using a 300 ms saturating actinic pulse at ~25,000 µmol photons m -2*s-1 and a Red LED matrix (Luxeon Rebel SM T High Power LED Red, LXM2 -PD01 -0050, Philips Lumiled, San Jose, California) to measure chlorophyll which was then captured by a CCD camera (AVT Manta 145 * M) equipped with a near infrared long pass filter (RT -830, Hoya Glass). Plants were imaged over three consecutive 24 * hr photoperiods ( Figure 2.2A, Supplemental Files 1 -9, timing and light intensities are described in Tables 2.5 -2.7). During the first day, the actinic light intensity remained constant at 100 µmol photons m -2*s-1 to collect growth chamber c onditions. Days two and three represented ramped lighting 47 perturbations. The photoperiod for day two was sinusoidal, beginning at 39 µmol photons m -2*s-1 and increasing in intensity by approximately 1.2 times every 30 * min until midday where it peaked at 50 0 µmol photons m -2*s-1, after which the light intensity decreased at the same rate every 30 * min. The photoperiod for day three was sinusoidal with brief fluctuations in light intensity. Starting at 39 µmol photons m -2*s-1, the light intensity was doubled after 15 * min followed by 1.5 -fold increase for 12 * min and the cycle repeated until peak intensities of 1000 and 500 µmol photons m -2*s-1 were cycled through at midday, after which the sinusoidal fluctuations decreased at the same rate as the increases. Ste ady state values for NPQ parameters of chlorophyll fluorescence were captured prior to the ramp to the next light intensity on days two and three, or hourly on day one and calculated as noted above for the fluorescence spectroscopy. Sequences of images wer e captured with a 60 ms delay between images for a 15 frame total for each measurement pre - during and post -saturation flash followed by images taken to correct for artifacts due to residual electrons in the CCD array. Images were analyzed using open sourc e software (ImageJ, NIH) modified in house to allow calculations of photosynthetic fluorescence parameters across selected regions of interest. 2.5.7 Photoinhibition of detached leaves. Plant leaves were excised and incubated in the dark for 3 * hr with the ir petioles submerged in either water or 3 * mM lincomycin to inhibit chloroplast protein translation (78). The leaves were then illuminated with red light at 1000 µmol photons m -2*s-1 using a red actinic light for indicated periods of time. During illuminat ion, the petioles of the leaves remained submerged in the treatment solution. Following illumination, leaves were allowed to dark adapt for 20 * min, after which the F 0 and F M values of chlorophyll fluorescence were measured in order to 48 determine F V/FM. For tobacco plants, leaf discs were soaked in a lincomycin solution and fluorescence parameters determined at 600 µmol photons m -2*s-1. To determine PSII activity, following photoinhibitory treatment, performed as above, leaves were dark adapted for 20 * min and then vacuum infiltrated with a 50 * µM DCMU solution. Analysis of PSII activity was determined from the amplitude of the ! A520 nm ECS signal using two saturating single -turnover flashes provided by a xenon lamp spaced 200 ms apart. The amplitude of the seco nd flash, corresponding to PSI centers capable of charge separation, was subtracted from the amplitude of the first flash, corresponding to both PSI and PSII centers capable of charge separation, to obtain the relative PSII photosystems capable of activity both before and after photoinhibition. Xenon flashes were judged to be fully saturating by ensuring that essentially identical results were obtained with a 50% weaker intensity. 2.5.8 Protein analysis. Photoinhibited leaf samples were collected as describ ed above and total leaf proteins were extracted as described in Livingston et al. (118). Analysis of chloroplast ATP synthase complexes was carried out on 20 * µg of protein using leaves collected from 3 Ð4 plants taken from the growth chamber. Analysis of Ps bA (D1) protein levels during photoinhibitory treatment was carried out on 30 * µg of total protein from 3 Ð4 leaves sampled during the photoinhibitory treatment time points as described above. Proteins were separated by SDS -PAGE and the ATPB (%-subunit) and PsbA (D1) proteins detected using commercially purchased antibodies (Agrisera, Vannas, Sweden). 2.5.9 1O2 detection. Plant leaves were excised and incubated in the dark for 3 * hr in 250 * µM Singlet Oxygen Sensor Green (SOSG, Life Technologies) prepared accor ding to manufacturerÕs instructions. 49 Petioles were maintained below the liquidÕs surface during the infiltration and were wrapped in a Kimwipe soaked in water during imaging to prevent drying. For valinomycin treatments, leaves were vacuum infiltrated with either SOSG solution or with SOSG supplemented with 50 * µM valinomycin and subsequently measured. Successful penetration of leaf epidermal and mesophyll cells, as well as SOSG penetrance throughout the cells was confirmed via confocal microscopy. The leave s were imaged in a chamber equipped with the same lighting as above. Qualitatively similar data were obtained for minira 3Ð1 leaves under both white and red (650 nm) LED illumination, ensuring that photosensitization of SOSG was not responsible for the sig nals obtained (122). Images were captured with a cooled CCD camera (AVT Bigeye G 132B -NIR) equipped with a 555 nm 10 nm band pass filter. Fluorescence excitation was provided via 458 nm LEDS (Cree Inc). Images were analyzed using ImageJ software. 2.5.10 Da ta analysis. All spectroscopic data were analyzed and figures generated using Origin 9.0 software (Microcal Software). Statistical analyses of data were performed in R package, utilizing two -way ANOVA to test for significant effects on photoinhibition (q I) from the interaction with either the ! pH or !! component of the pmf . 2.6 Acknowledgements. This work was supported by the U.S. Department of Energy (DOE), Office of Science, Basic Energy Sciences (BES) under Award number DE -FG02 -91ER20021 and the MSU Center for Advanced Algal and Plant Phenotyping (CAAPP). AWR was supported by a Biotechnology and Biological Sciences Research Council (BBSRC) grant (BB/K002627/1) and the Royal Society Wolfson Research Merit Award. 50 Supplemental Figure 2.1: Whole plant fluorescence imaging phenotyping of minira 3Ð1 mutant. Three week old plants were imaged over three consecutive 16 -hr photoperiods and fluorescent measurements taken at the end of each light transition for LEF (A ÐC), qE (DÐF), and q I (GÐI) for Ws-2 and minira 3Ð1. Values represent mean of n ( 3 ± s.d., all of which were imaged in the same experiment. Timing and illumination are the same as Figure 2.2 and are detailed in Tables 2.5-2.7. 51 Supplemental Figure 2.2: Increased pH -dependent quenching co rrelates with increased photoinhibitory quenching. Whole plant fluorescent phenotypes were measured over three consecutive photoperiods. Day one consisted of a single irradiance level (open symbols), day two of sinusoidal irradiance (half filled symbols), and day three of sinusoidal irradiance interrupted by bright fluctuations (closed symbols). The pH -dependent fluorescence quenching (q E) and photoinhibitory quenching (q I) were integrated over time for each plant for each photoperiod to determine how prolo nged exposure to an increased pmf influences the extent of photoinhibition. Data represent the integrated total for each day (n ( 3, ± s.d). 52 Supplemental Figure 2.3: Whole plant fluorescence imaging phenotyping of minira 11Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 53 Supplemental Figure 2.4: Whole plant fluorescence imaging phenotyping of minira 14Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 54 Supplemental Figure 2.5: Whole plant fluorescence imaging phenotyping of minira 12Ð2 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 55 Supplemental Figure 2.6: Whole plant fluorescence imaging phenotyping of minira 8Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 56 Supplemental Figure 2.7: Whole plant fluorescence imaging phenotyping of minira 6Ð2 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 57 Supplemental Figure 2. 8: Whole plant fluorescence imaging phenotyping of minira 4Ð2 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 58 Supplemental Figure 2.9: Whole plant fluorescence imaging phenotyping of minira 6Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 59 Supplemental Figure 2.10: Whole plant fluorescence imaging phenotyping of minira 7Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 60 Supplemental Figure 2 .11: Whole plant fluorescence imaging phenotyping of minira 3Ð2 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 61 Supplemental Figure 2.12: Whole plant fluorescence imaging phenotyping of minira 4Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 62 Supplemental Figure 2.13: Whole plant fluorescence imaging phenotyping of minira 9Ð1 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 63 Supplemental Figure 2 .14: Whole plant fluorescence imaging phenotyping of minira 4Ð3 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 64 Supplemental Figure 2.15: Whole plant fluorescence imaging phenotyping of minira 12Ð3 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 65 Supplemental Figure 2.16: Whole plant fluorescence imaging phenotyping of minira 2Ð2 mutant. For details, please refer to the legend of Supplemental Figure 2.1 . 66 Supplemental Figure 2.17: Reduction kinetics of P700 +. Light -dependent P700+ reduction half -times (mean ± s.d., n = 3) of wild type, minira 3Ð1 and minira 14Ð1 (A). Quenching of the 810 nm absorbance signal was followed during a brief dark interval and the half time of the first order decay determined at each light intensity. 67 Supplemental Figure 2.18: The electric field component of the pmf dominates under high pmf conditions. ECS measurements were performed to determine the partitioning of the light -driven pmf between ! pH and !!. Measurements were performed at 100 (solid symbols), 300 (half filled symbols), and 500 µmol photons m '2s'1 (open symbols) and the total pmf (ECS t) as well as the composition of the pmf determined as in Figure 2.5 . As the total pmf increases, the fraction of pmf stored as ECS inv (proportional to ! pH) (A) decreases linearly and is observed in all lines regardless of mutation, while the electric field (ECS ss) (proportional to !!) (B) increases linearly with total pmf , becoming a large fraction of the total pmf under high pmf conditions. Symbols represent the same plant for each light intensity (mean ± s.d., n = 3). ECS units were defined as the deconvoluted !A520 µg chlorophyll '1 cm2. 68 Supplemental Figure 2.19: Tobacco ATPC1 antisense knockdown increase !! partitioning under high pmf conditions. The partitioning of the pmf in wild type Samsun (black) and ATPC1 (red) "-subunit antisense line were determined from the deconvoluted ECS signal at ! A520 nm (A). Following the light -dark transition (time 0 s), the ECS amplitude drop, which is proportional to the total pmf , is larger in the ATPC1 line due to the substantial knockdown o f ATP synthase complexes and an inability to efflux protons from the lumen. The inversion of the ECS signal during the dark period, proportional to the ! pH, represents a larger fraction of the pmf in the wild type than in ATPC1, indicating an increase in !! storage in the higher pmf ATPC1. Light conditions are represented above the traces and indicate the light (yellow) and dark (black) intervals. Leaf discs from another ATPC1 plant and Samsun were subjected to 600 µmol photons m '2s'1 light for the indicate d times in the presence of water (open symbols) or lincomycin (closed symbols). Increases in q I are more rapid in the higher pmf ATPC1 line when PSII repair is blocked with lincomycin. 69 Supplemental Figure 2.20: Uncoupling !! decreases SOSG fluorescence in minira 3Ð1. Minira 3Ð1 leaves were vacuum infiltrated with either SOSG (solid triangles) or SOSG and 50 µM valinomycin (crossed triangles) to decrease the photosynthetic !!. Leaves were illuminated at a constant 100 µmol m '2s'1 intensity and SOSG fluor escence detected for 60 min. 70 Supplemental Figure 2.21: Fluctuations in light intensity result in transient ECS spikes. Wild type plants were measured under fluctuating light (A) and the ECS measurements taken 10 s after each intensity fluctuation from lower to higher light (A, open squares). The resulting deconvoluted ! A520nm ECS signals show rapid, transient ECS 'spikes' induced by rapid !! transients before the ! pH component can be altered (B). The upward spike represents the effect of rapidly increa sing the light intensity. The downward spikes seen towards the ends of the traces reflect the transients that occur when the actinic light is switched to the lower light intensity at the end of the fluctuation. A rough estimate of the extent of the !, impo sed by the spikes can be obtained by comparing the ECS signals with the calibration and results presented in Takizawa et al. (47), which estimated that the !! imposed by a saturating, single -turnover flash to thylakoids was about 40 mV. Based on this calibration value, the extent of basal pmf formed by equilibration with ATP (i.e. the dark pmf level or pmf d) was estimated to be about 112 mV (Takizawa et al., 20 07), 60 mV of which is stored in !! if the partitioning is 0.5. From the dark -interval relaxation kinetics of ECS under steady -state conditions (see Figure 2.7B and Supplemental Figure 2.17), we then expect an additional light -driven pmf under steady -state conditions to range from 150 Ð200 mV, and given that the fraction of this pmf stored as !! ranged from 0.20 Ð0.60, we estimate a range for steady -state light -driven !! between 30 and 120 mV. The transient spikes in !! generated during light fluctuations are likely to be essentially all stored in !! (see main text) and range in amplitude between 60 Ð80 mV, so that the highest amplitude !, imposed during these conditions likely falls in the broad range between 150 Ð260 mV. ECS units were defined as the deconvolu ted ! A520 µg chlorophyll '1 cm2. 71 Table 2.1: Oligonucleotide sequences utilized for adapter ligation mutagenesis. Mutation minira # Adapto r Oligonucleotide sequences with mutated nucleotide underlined and in bold I201V minira 4 DMP 27 5Õ GATCTGTGACGTTAATGGAACCTGTGTGGATGCTGCGGAAGATGAGTTTTTCAGGTT 3Õ DMP 28 5Õ AACCTGAAAAACTCATCTTCCGCAGCATCCACACAGGTTCCATTAACGTCACA 3Õ N202K minira 5 DMP 29 5Õ GATCTGTGACATTAAAGGAACCTGTGTGGATGCTGCGGAAGATGAGTTTTTCAGGTT 3Õ DMP 30 5Õ AACCTGAAAAACTCATCTTCCGCAGCATCCACACAGGTTCCTTTAATGTCACA 3Õ A209I minira 6 DMP 31 5Õ GATCTGTGACATTAATGGAACCTGTGTGGATGCTATCGAAGATGAGTTTTTCAGGTT 3Õ DMP 32 5Õ AACCTGAAAAACTCATCTTCGATAGCATCCACACAGGTTCCATTAATGTCACA 3Õ F213M minira 7 DMP 33 5Õ GATCTGTGACATTAATGGAACCTGTGTGGATGCTGCGGAAGATGAGATGTTCAGGTT 3Õ DMP 34 5Õ AACCTGAACATCTCATCTTCCGCAGCATCCACACAGGTTCCATTAATGTCACA 3Õ 72 Table 2.2 : Oligonucleotide sequences utilized for adapter ligation mutagenesis to introduce secondary mutations. Mutation minira # Adapto r Oligonucleotide sequences with mutated nucleotide underlined and in bold P194M minira 2 DMP 23 5Õ GTACACAAAGTTTGTCTCTTTGGTCAAATCAGAACCCGTGATCCACACGCTACTGCCTTTATCAATGAAAGGAGA 3Õ DMP 24 5Õ GATCTCTCCTTTCATTGATAAAGGCAGTAGCGTGTGGATCACGGGTTCTGATTTGACCAAAGAGACAAACTTTGT 3Õ E183D minira 1 DMP 21 5Õ GTACACAAAGTTTGTCTCTTTGGTCAAATCAGATCCCGTGATCCACACGCTACTGCCTTTATCACCTAAAGGAGA 3Õ DMP 22 5Õ GATCTCTCCTTTAGGTGATAAAGGCAGTAGCGTGTGGATCACGGGATCTGATTTGACCAAAGAGACAAACTTTGT 3Õ 73 Table 2.3: Oligonucleotide sequences utilized for splicing by overlap extension PCR. Mutation minira # Region Primer Primer sequence T218S minira 8 Fragment 1 DMP49 5Õ AACTGTCAATTTCCCTTCTTTACTCGTTAACCT 3Õ DMP 45 5Õ TCCTGCAGCCCGGGAACAAAAAAAT 3Õ Fragment 2 DMP 50 5Õ TGAGTTTTTCAGGTTAACGAGTAAAGAAGGG 3Õ DMP 46 5Õ GCGGCCGCTCTAGACAAATCAAAC 3Õ E220D minira 9 Fragment 1 DMP51 5Õ AACTGTCAATTTCCCGTCTTTTGTCGTTAAC 3Õ DMP 45 5Õ TCCTGCAGCCCGGGAACAAAAAAAT 3Õ Fragment 2 DMP 52 5Õ CAGGTTAACGACAAAAGACGGGAAATT 3Õ DMP 46 5Õ GCGGCCGCTCTAGACAAATCAAAC 3Õ T224A minira 10 Fragment 1 DMP53 5Õ GTCTCTCTTTCAACTGCCAATTTCCC 3Õ DMP 45 5Õ TCCTGCAGCCCGGGAACAAAAAAAT 3Õ Fragment 2 DMP 54 5Õ CGACAAAAGAAGGGAAATTGGCAGTTGA 3Õ DMP 46 5Õ GCGGCCGCTCTAGACAAATCAAAC 3Õ E228T minira 11 Fragment 1 DMP55 5Õ TGTTGGTGTCCTAAAAGTCGTTCTTTCAACTGT 3Õ DMP 45 5Õ TCCTGCAGCCCGGGAACAAAAAAAT 3Õ Fragment 2 DMP 56 5Õ GAAATTGACAGTTGAAAGAACGACTTTTAGGA 3Õ DMP 46 5Õ GCGGCCGCTCTAGACAAATCAAAC 3Õ I201V- N202K minira 12 Fragment 1 DMP57 5Õ CACACAGGTTCCTTTCACGTCACAGATCTC 3Õ DMP 45 5Õ TCCTGCAGCCCGGGAACAAAAAAAT 3Õ Fragment 2 DMP 58 5Õ GAGATCTGTGACGTGAAAGGAACCTGTGTG 3Õ DMP46 5Õ GCGGCCGCTCTAGACAAATCAAAC 3Õ 74 Table 2.4: Synthetic gene constructs incorporating multiple ATPC2 mutations into ATPC1. Domain minira # Synthetic gene fragment sequence with the new domain in bold. Restriction enzyme sites are underlined. 194-241 minira 16 aattaaTGTACACAAAGTTTGTCTCTTTGGTCAAATCAGAACCCGTGATCCACACGCTACTGCCTTTATCGATGAAAGGAGAGTCTTGTGATGTGAAAGGTGAGTGTGTTGATGCTATCGAGGATGAGATGTTTAGGCTAACGAGCAAAGATGGGAAGTTAGCTGTGGAAAGGACCAAGCTTGAAGTTGAGAAGCCTGAGATCTCACCGTTGATGCAATTCGAGCAAGACCCTGTTCAGATTCTTGATGCTTTGTTGCCTCTGTATCTTAACAGTCAGATTCTTAGGGCATTACAGGAGTCATTGGCTAGTGAGCTTGCAGCTAGAATGAGTGCAATGAGTAGTGCTTCGGATAATGCATCGGATCTCAAGAAATCGCTTTCGATGGTGTATAATAGAAAGCGTCAAGCTAAGATTACTGGAGAGATTCTTGAGATTGTTGCTGGAGCTAATGCACAGGTTTGATTTGTCTAGAttaatt 213-228 minira 14 aattaaTGTACACAAAGTTTGTCTCTTTGGTCAAATCAGAACCCGTGATCCACACGCTACTGCCTTTATCACCTAAAGGAGAGATCTGTGACATTAATGGAACCTGTGTGGATGCTGCGGAAGATGAGATGTTTAGGCTAACGAGCAAAGATGGGAAGTTAGCTGTGGAAAGGACCACTTTTAGGACACCAACAGCTGATTTCTCGCCGATCTTGCAATTCGAGCAAGACCCTGTTCAGATTCTTGATGCTTTGTTGCCTCTGTATCTTAACAGTCAGATTCTTAGGGCATTACAGGAGTCATTGGCTAGTGAGCTTGCAGCTAGAATGAGTGCAATGAGTAGTGCTTCGGATAATGCATCGGATCTCAAGAAATCGCTTTCGATGGTGTATAATAGAAAGCGTCAAGCTAAGATTACTGGAGAGATTCTTGAGATTGTTGCTGGAGCTAATGCACAGGTTTGATTTGTCTAGAttaatt I198S minira 3 aattaaTGTACACAAAGTTTGTCTCTTTGGTCAAATCAGAACCCGTGATCCACACGCTACTGCCTTTATCACCTAAAGGAGAGAGCTGTGACATTAATGGAACCTGTGTGGATGCTGCGGAAGATGAGTTTTTCAGGTTAACGACAAAAGAAGGGAAATTGACAGTTGAAAGAGAGACTTTTAGGACACCAACAGCTGATTTCTCGCCGATCTTGCAATTCGAGCAAGACCCTGTTCAGATTCTTGATGCTTTGTTGCCTCTGTATCTTAACAGTCAGATTCTTAGGGCATTACAGGAGTCATTGGCTAGTGAGCTTGCAGCTAGAATGAGTGCAATGAGTAGTGCTTCGGATAATGCATCGGATCTCAAGAAATCGCTTTCGATGGTGTATAATAGAAAGCGTCAAGCTAAGATTACTGGAGAGATTCTTGAGATTGTTGCTGGAGCTAATGCACAGGTTTGATTTGTCTAGAttaatt 75 Table 2.5 : Timing and light profile of imaging day 1 . Light Intensity Day 1 Duration at Intensity (min) Time of Day 0 359.4 6:00 100 58 6:58 0 2 6:00 100 58 7:58 0 2 7:00 100 58 8:58 0 2 8:00 100 58 9:58 0 2 9:00 100 58 10:58 0 2 10:00 100 58 11:58 0 2 11:00 100 58 12:58 0 2 12:00 100 58 13:58 0 2 13:00 100 58 14:58 0 2 14:00 100 58 15:58 0 2 15:00 100 58 16:58 0 2 16:00 100 58 17:58 0 2 17:00 100 58 18:58 0 2 18:00 100 58 19:58 0 2 19:00 100 58 20:58 0 2 20:00 100 58 21:58 0 2 22:00 76 Table 2.6: Timing and light profile of imaging day 2 . Light Intensity Day 2 Duration at Intensity (min) Time of Day 0 359.4 6:00 39 28 6:28 0 2 6:30 80 28 6:58 0 2 7:00 123 28 7:28 0 2 7:30 167 28 7:58 0 2 8:00 210 28 8:28 0 2 8:30 253 28 8:58 0 2 9:00 294 28 9:28 0 2 9:30 333 28 9:58 0 2 10:00 370 28 10:28 0 2 10:30 402 28 10:58 0 2 11:00 431 28 11:28 0 2 11:30 455 28 11:58 0 2 12:00 475 28 12:28 0 2 12:30 489 28 12:58 0 2 13:00 497 28 13:28 0 2 13:30 500 28 13:58 0 2 14:00 500 28 14:28 0 2 14:30 497 28 14:58 0 2 15:00 489 28 15:28 77 Table 2.6 (continued ): 0 2 15:30 0 2 15:30 475 28 15:58 0 2 16:00 455 28 16:28 0 2 16:30 431 28 16:58 0 2 17:00 402 28 17:28 0 2 17:30 370 28 17:58 0 2 18:00 333 28 18:28 0 2 18:30 294 28 18:58 0 2 19:00 253 28 19:28 0 2 19:30 210 28 19:58 0 2 20:00 167 28 20:28 0 2 20:30 123 28 20:58 0 2 21:00 80 28 21:28 0 2 21:30 39 28 21:58 0 2 22:00 78 Table 2.7: Timing and light profile of imaging day 3 . Light Intensity Day 3 Duration at Intensity (min) Time of Day 0 359.4 6:00 39 18 6:18 0 2 6:20 78 8 6:28 0 2 6:30 80 18 6:48 0 2 6:50 161 8 6:58 0 2 7:00 123 18 7:18 0 2 7:20 246 8 7:28 0 2 7:30 167 18 7:48 0 2 7:50 333 8 7:58 0 2 8:00 210 18 8:18 0 2 8:20 420 8 8:28 0 2 8:30 253 18 8:48 0 2 8:50 506 8 8:58 0 2 9:00 294 18 9:18 0 2 9:20 588 8 9:28 0 2 9:30 333 18 9:48 0 2 9:50 667 8 9:58 0 2 10:00 370 18 10:18 0 2 10:20 739 8 10:28 0 2 10:30 402 18 10:48 79 Table 2.7 (continued ): 0 2 10:50 805 8 10:58 0 2 11:00 431 18 11:18 0 2 11:20 862 8 11:28 0 2 11:30 455 18 11:48 0 2 11:50 911 8 11:58 0 2 12:00 475 18 12:18 0 2 12:20 949 8 12:28 0 2 12:30 489 18 12:48 0 2 12:50 977 8 12:58 0 2 13:00 497 18 13:18 0 2 13:20 994 8 13:28 0 2 13:30 500 18 13:48 0 2 13:50 1000 8 13:58 0 2 14:00 500 18 14:18 0 2 14:20 1000 8 14:28 0 2 14:30 497 18 14:48 0 2 14:50 994 8 14:58 0 2 15:00 489 18 15:18 0 2 15:20 977 8 15:28 0 2 15:30 475 18 15:48 80 Table 2.7 (continued ): 0 2 15:50 949 8 15:58 0 2 16:00 455 18 16:18 0 2 16:20 911 8 16:28 0 2 16:30 431 18 16:48 0 2 16:50 862 8 16:58 0 2 17:00 402 18 17:18 0 2 17:20 805 8 17:28 0 2 17:30 370 18 17:48 0 2 17:50 739 8 17:58 0 2 18:00 333 18 18:18 0 2 18:20 667 8 18:28 0 2 18:30 294 18 18:48 0 2 18:50 588 8 18:58 0 2 19:00 253 18 19:18 0 2 19:20 506 8 19:28 0 2 19:30 210 18 19:48 0 2 19:50 420 8 19:58 0 2 20:00 167 18 20:18 0 2 20:20 333 8 20:28 0 2 20:30 123 18 20:48 81 Table 2.7 (continued ): 0 2 20:50 246 8 20:58 0 2 21:00 80 18 21:18 0 2 21:20 161 8 21:28 0 2 21:30 39 18 21:48 0 2 21:50 78 8 21:58 0 2 22:00 82 Table 2. 8: Chlorophyll content of wild type (Ws -2) and minira leaves. Genotype Chlorophyll ( µg cm-2) Ws-2 16.55 + 0.51 minira 2-2 13.00 + 0.17* minira 3-1 8.24 + 1.62* minira 3-2 13.04 + 1.45 minira 4-1 14.55 + 1.18 minira 4-2 14.05 + 1.34 minira 4-3 14.99 + 0.78 minira 6-1 13.64 + 0.78* minira 6-2 13.74 + 0.85* minira 7-1 13.48 + 0.25* minira 8-1 10.51 + 1.53* minira 9-1 12.53 + 2.85 minira 11-1 11.98 + 1.52* minira 12-2 8.46 + 2.28* minira 12-3 12.28 + 0.94* minira 14-1 13.87 + 1.65 83 Chapter 3 Hacking the thylakoid proton motive force for improved photosynthesis: modulating ion flux rates that control proton motive force partitioning into !! and !pH 2 Geoffry A. Davis, A. William Rutherford, David M. Kramer 2 This ch apter was published as: Davis, GA, Rutherford, AW, & Kramer, DM. 2017 Hacking the thylakoid proton motive force for improved photosynthesis: modulating ion flux rates that control proton motive force partitioning into !! and !pH. Philos Trans R Soc Lond B Biol Sci, 372(1730). doi: 10.1098/rstb.2016.0381 84 3.1 Abstract There is considerable interest in improving plant productivity by altering the dynamic responses of photosynthesis in tune with natural conditions. This is exemplified by the Ôenergy -dependent' form of non -photochemical quenching (q E), the formation and decay of which can be considerably slower than natural light fluctuations, limiting photochemical yield. In addition, we recently reported that rapidly fluctuating light can produce field recombination -induced photodamage (FRIP), where large spikes in electric field across the thylakoid membrane ( !!) induce photosystem II recombination reactions that produce damaging singlet oxygen ( 1O2). Both q E and FRIP are directly linked to the thylakoid proton motive force ( pmf ), and in particular, the slow kinetics of partitioning pmf into its ! pH and !! components. Using a series of computational simulations, we explored the possibility of Ôhac king' pmf partitioning as a target for improving photosynthesis. Under a range of illumination conditions, increasing the rate of counter -ion fluxes across the thylakoid membrane should lead to more rapid dissipation of !! and formation of ! pH. This would result in increased rates for the formation and decay of q E while resulting in a more rapid decline in the amplitudes of !!-spikes and decreasing 1O2 production. These results suggest that ion fluxes may be a viable target for plant breeding or engineering . However, these changes also induce transient, but substantial mismatches in the ATP:NADPH output ratio as well as in the osmotic balance between the lumen and stroma, either of which may explain why evolution has not already accelerated thylakoid ion flu xes. Overall, though the model is simplified, it recapitulates many of the responses seen in vivo , while spotlighting critical aspects of the complex interactions between pmf components and photosynthetic processes. By making the programme available, we hope to enable the community of photosynthesis researchers to further explore and test specific hypotheses. 85 3.2 Introduction This opinion/hypothesis paper was inspired by the r ecent Royal Society symposium on ÔEnhancing photosynthesis in crop plants: targets for improvement' (http://www.rsc.org/events/download/Document/cee7d4f2 -9ff1 -477b-b155-3e2492577d77) that brought together experts in a range of photosynthetic processes. A p rominent theme of several of the presentations was the sensitivity of photosynthesis to rapid, rather than gradual, changes in environmental conditions. Of particular interest was the kinetic mismatch between fluctuations in photosynthetically active radia tion (PAR), which can change by orders of magnitude within a second, and the relatively slow onset of photoprotective mechanisms (123, 124). Indeed, there is growing evidence that this mismatch can sensitize both photosystem I (PSI) and photosystem II (PSI I) to oxidative photodamage (36, 97). The focus of the present paper is the irreversible damage to PSII (to D1 and other subunits) due to singlet oxygen ( 1O2) generated by PSII charge recombination. It has also been proposed that the slow reversal of photo protection mechanisms can lead to loss of photochemical productivity when light levels are suddenly decreased (125, 126). Thus, such kinetic mismatches appear to be good engineering targets for increasing the efficiency and resilience of photosynthesis. He re, we present an extended re -examination of one of the key processes that controls and regulates photosynthesis, the thylakoid proton motive force (pmf ), its components, and some emerging effects on photosynthetic reaction centres. The energy -storing proc esses of photosynthesis start with the capture of PAR by photoactive pigments, transfer of the energy to specialized chlorophyll molecules in PSI and PSII, inducing the transfer of electrons through a series of redox intermediates to ultimately generate NA DPH from NADP + (127). The electron transfer steps are tightly coupled to proton transfer reactions into the thylakoid lumen, storing potential energy in the pmf to drive the 86 synthesis of ATP (61, 76). Both NADPH and ATP, in the correct ratios, are required for driving the assimilation of CO 2 and other cellular processes (121). The electron and proton transfer processes can be highly efficient, but when energy capture outpaces the capacity of photosynthesis Ña situation that can occur at high light and/or und er adverse environmental conditions Ñreactive intermediates can accumulate in the photosynthetic apparatus, leading to generation of reactive oxygen species (ROS), mainly O 2-¥ in PSI and mainly 1O2 in PSII, and these are responsible for oxidative photodamag e. A range of photoprotective mechanisms have evolved to ameliorate photodamage and its effects, including non -photochemical quenching (NPQ) processes such as the qE response (6), a complex cycle to repair damaged PSII (128), chloroplast movements (129), cyclic electron flow (130, 131), redox tuning to redirect back -reactions to non -ROS producing pathways (56, 132) and alternative electron acceptor systems (5, 133). Despite the diversity and complexity of these processes, in general, they result in the loss of light energy, for example, the decreased efficiency of light capture incurred by activation of NPQ (134), charge recombination (56) or the dissipation of redox energy when electrons are passed to the flavodiiron O 2 reductases (123). Consequently, photo synthetic organisms appear to be constantly balancing the trade -offs between efficient photochemistry and the avoidance of toxic side reactions. In higher plants and green algae, the pmf plays a central role in regulating key photoprotective mechanisms, by responding to changes in both energy input and the physiological status of the chloroplast (61). It is, therefore, worthwhile to review the biophysical properties of the photosynthetic machinery that controls the partitioning of pmf into !! and ! pH over different time -scales. In thermodynamic terms, the pmf can be described as the sum of two driving forces: 87 !"# !!!!!!!!!"!!!" (Eq. 1) where !! and ! pH represent the differences in electric field, expression difference in volts, and pH, respectively, between the lumenal and stromal faces of the thylakoid membrane, R is the universal gas constant and F is Faraday's constant. Over a broad ra nge of physiological conditions, !! and ! pH appear to be thermodynamically and kinetically equivalent drivers of the ATP synthase (3, 4). On the other hand, storing pmf in !! and ! pH has distinct impacts on cellular processes. Most notably, storing energy in ! pH imposes a substantial change in pH in one or more cellular compartments. In mitochondria, pmf is held mainly as !!, allowing enzymes to operate at optimal pH ranges. In chloroplasts, the build -up of ! pH results in acidification of the thylakoid lume n, which acts to feedback regulate (or control) critical steps in the light reactions (reviewed in 5), including (i) the activation of the photoprotective q E response (through activation of violaxanthin deepoxidase and protonation of PsbS (6)); and (ii) Ôphotosynthetic control' of electron flow at the cytochrome b6f complex (reviewed in 13), preventing the accumulation of electrons on PSI that would otherwise lead to severe PSI photodamage (5, 47, 123, 124). There have also been proposals that PSII can be regulated or inhibited (13, 15) at low lumen pH, for example, by acid -induced release of Ca 2+ from the oxygen -evolving complex (OEC), or by limiting electron flow by slowing of the OEC S -state transitions (15, 63). Early work on isolated thyl akoids suggested that pmf was stored mainly as ! pH, but more recent work suggests that a pure ! pH pmf is incompatible with the known pH dependencies of photosynthetic processes (13, 39). A range of in vivo studies (5, 36, 47, 70, 135-138) support the view that the pmf is actively partitioned into !! and ! pH components to avoid severe 88 restrictions on b6f activity or acid -induced damage to lumenal components, while balancing the needs for efficient energy storage and activation of lumen pH -responsive photopro tective processes (39). These arguments are supported by our simulations (below) and straightforward thermodynamic considerations, which indicate that: with ! GATP (the free energy of hydrolysis of ATP) between 40 and 45 kJ mol -1, a stromal pH of 7.8, and t he coupling stoichiometry for protons/ATP, n, of 4.67, the maximal lumen pH (even before illumination) should range between 6 and 6.5 units, near or below the pKa values that govern the activation of q E and the control of cytochrome b6f activity. In short, with 100% ! pH, the photosynthetic electron transport chain should be strongly downregulated even at low light. Nevertheless, there are opposing views that maintain pmf is stored almost exclusively in ! pH under steady -state conditions (139). It is worthwhile to note, however, that the phenomena discussed in this review are associated more with the dynamics of !! and ! pH rather than their steady -state values. In any case, the balancing of !!/ ! pH, and its kinetics, are dependent on the re gulation of counter -ion homeostasis in the chloroplast (2), and recent work from several laboratories has identified putative components of these ion homeostatic machineries, including a thylakoid potassium channel (27), a K +/H+ antiporter (28), as well as transporters for other charged species (29-32, 140). There are also indications that the !!/ ! pH balance is controlled by the synthesis of membrane -permeable weak bases, such as putrescene, that effectively increase the proton buffering capacity of the lum en (54). 3.3 Interactions of the pmf with PSII: Importance for energy storage and photodamage. The photosystems can be viewed as Ôenergy traps' that capture energy from sunlight in the form of quasi -stable charge -separated states. To achieve this role, ev olution has tuned the redox properties of reaction centre cofactors so that each progressive electron transfer reaction 89 occurs more rapidly than the decay by other routes (e.g. back -reactions or charge recombination), successively stabilizing the resulting charge -separated states and minimizing the losses to back -reactions or recombination to the point where quantum efficiency is near unity but at the cost of free energy losses at each step (56). Despite the high quantum yield of formation of stable charge -separated states, electron transfer back -reactions and charge recombination can occur, the rates of which depend on the energetics of the free energy gap, reorganizational energies and donorÐacceptor distances between the oxidized and reduced components in the reaction centres (56, 141, 142). For PSII, the most important recombination reaction (both in terms of rates and consequences for photodamage) occurs from the P 680+Pheo - radical pair state (with P 680 oxidized and pheophytin reduced), which can be formed via both initial forward electron transfer, or by back -reactions via thermal activation of ÔstableÕ intermediates in the PSII photocycle, e.g. S 2QB-, the state formed when dark -adapted PSII undergoes a single photochemical reaction. Further thermal activation of P 680+Pheo - can repopulate the P * state, leading to essentially the full reversal of the initial light reactions, resulting in the emission of Ôdelayed fluorescence' (143), though owing to the hi gh energy of activation this process has a low quantum efficiency (143). The intensity and temperature dependence of delayed fluorescence have been extensively used to estimate the energetics of reaction centres, and are potentially important probes for th e more deleterious processes discussed below. More importantly for this discussion, the P +680Pheo - state can also decay non -radiatively, either directly to ground state, or via the triplet state of P 680 (3P680) (144). In turn, 3P680 can interact with molec ular O 2 to form singlet O 2 (1O2), a highly ROS that can damage both PSII and other cellular components (93). The rates and yield of back -reactions leading to P 680+Pheo - recombination should be accelerated under any conditions that make the energy gaps betw een P 680+Pheo - and the 90 subsequent radical pairs (e.g. P 680+QA-) shallower. For example, it is well known that when plants are treated with herbicides that are Q B site inhibitors, such as 3 -(3,4-dichlorophenyl) -1,1-dimethylurea (DCMU) or atrazine, which blo ck electron transfer from Q A- to Q B, the recombination reactions speed up by about 10 - to 20 -fold (99, 145), and the enhanced formation of 1O2 triggers processes that can damage the cell. Our recent work describes a more physiological process that we term field recombination -induced photodamage (FRIP) in which large spikes in the photosynthetic !!, such as those caused by rapid fluctuations in light intensity, destabilize the PSII photochemically generated charge pairs, accelerating PSII back -reactions, charge recombination and 1O2 production (36). Our results suggest that FRIP represents both a li mitation for energy storage as well as a limitation to productivity from losses due to photodamage, especially under rapidly fluctuating conditions found in nature. 3.4 Computational exploration of the effects of altering pmf storage kinetics on q E and fie ld recombination Ðinduced photodamage. NPQ and FRIP should both be sensitive to kinetic mismatches between the generation of !! and ! pH and subsequent regulatory responses, and it is conceivable that engineering these processes could lead to improved photos ynthesis. To test this possibility, we updated a previously published computational model for pmf (2, 146) that describes how the intrinsic biophysical properties of the thylakoid system impact the extent and kinetics of pmf storage in !! and ! pH, and how these properties may affect the susceptibility of photosynthesis to photodamage under rapidly fluctuating conditions. While simplified, our model recapitulates many key features of the pmf . The biophysical bases of the current model are similar to those presented in other models (147), and thus using our parameters, one would expect similar results. The updated model includes the effects of !!, ! pH and Q A redox state on FRIP. The code was 91 written in Python 3.5 using open source modules and is presented in t he form of a detailed Jupyter (www.jupyter.org) notebook, which is included in the Appendix and freely available online at Github (www.github.com/protonzilla/Delta_Psi_Py), allowing the reader to download, modify, extend and explore variations of the simul ations presented here. The details of the code and references to the parameter set are presented in the extensive annotation in the code and accompanying explanatory notes in the Appendix and on the Github site. 3.5 The effects of !pH and !! on PSII recom bination and 1O2 production in vivo . As illustrated in figure 3.1a, the cofactors in PSII are situated such that charge separation reactions occur across the low dielectric of the thylakoid membrane, and thus the movement of electrons from P 680 (near the luminal face), through pheophytin (Pheo) and to Q A (near the stromal face) directly contribute to thylakoid !!. In addition, the oxidation of water at the OEC deposits protons into the lumen, while the reduction of plastoquinone (PQ) at the Q B site takes up protons from the stroma, contributing to ! pH. It follows that Ôbackpressure' from either of the pmf components could accelerate recombination reactions, as has been amply demonstrated in past work (38, 148). However, !! and ! pH should have dif ferential effects on recombination depending on the properties of the charge pair involved. The !! component will primarily affect the energetics of Ôelectrogenic' electron transfer reactions, i.e. those that occur vectorially across the thylakoid membrane , by shifting the equilibrium constant for these reactions towards the electron carrier closer to the lumenal side of the membrane (i.e. the positively charged side; figure 3.1b,c). The effect of !! on charge -separated states is dependent upon the physical distance between the redox cofactors and the perpendicular distance from the membrane edge, i.e. electron transfer between cofactors within the thylakoid membrane becomes progressively more destabilized by !! as the distance between charge -stable states s pans larger distances 92 across the membrane. For example, electron transfer from Pheo - to Q A moves a charge about halfway across the thylakoid dielectric, so that adding a !! of 120 mV should shift the free energy drop for this this reaction by about 60 meV, and alter the equilibrium constant for sharing the electron by a factor of about 10. Figure 3.1 : Photosynthetic electron transfer energetics are influenced by membrane orientation and membrane potential. (a) PSII cofactors are oriented within the complex so that light -driven forward electron transfer induces the formation of a trans -thylakoid !!. Subsequent proton uptake (red arrows) at Q B and release at the oxygen -evolving complex will contribute to the ! pH. During linear electron flow, additional !! (blue arrows) is generated by light -induced charge separation in photosystem I, while both !! and ! pH are generated by the Q -cycle at the cytochrome b6f complex. (b) The loss of free energy during PSII forwar d electron transfer energetically stabilizes (thick lines) the charge -separated state, impeding (thin lines) recombination. (c) Imposing either !! or ! pH decreases this stabilization energy, increasing the rate of recombination, leading to the generation of 3P680, which can interact with O 2 to form the toxic 1O2 species. The ! pH component, on the other hand, should primarily affect recombination reactions that involve the uptake or deposition of protons into the aqueous compartments. The pH of the 93 stroma i s thought to be relatively constant in the light, ranging between 7.0 in the dark to 8.0 in the light (149, 150). This pH change has a differential effect on Q A- versus Q B-, which are both located on the stromal side of the protein, during the initial dark Ðlight transition in stromal pH. QA- is unaffected by the light -induced alkalization of the stroma because no protonation reactions are involved in its redox chemistry. For Q B-, however, a proton is taken up when it is formed, and so a proton must be r eleased when it is oxidized, thus favouring re -oxidation of Q B- with increasing stromal pH and destabilization of the semiquinone (151). This affect, however, is likely to remain constant during the course of the day in chloroplasts, as the stromal pH appe ars relatively stable. On the luminal side, a decreased lumen pH via proton transfer reactions will destabilize the OEC S -states if the redox transition involves proton release. Past work (16) suggests that, over the relevant lumenal pH range (approx. from 5.5 to 7.5), the proton release pattern is 1, 0, 1, 2 for transitions S0 - S1, S1 - S2, S2 - S3 and S3 - S4 - S0. The respective back -reactions should involve proton uptake by the OEC, and thus will be destabilized by low lumen pH. The S0 - S1 and S3 - S4 - S0 transitions are considered to be irreversible, so once formed should not be prone to recombination. S2 and S3 recombine with Q A- and Q B- (with the back -reaction via P +680Pheo - as the dominant pathway); however, because the S2 - S1 back -reaction does not involve a proton transfer its rate should be unaffected by lowering the lumen pH. Therefore, only PSII centres in the S3 state should display pH -dependent increases in recombination rates due to lowering the lumen pH (by 10 -fold for a decrease of 1 pH unit). By contrast, !! should affect recombination for both S2 and S3 states. Under continuous light, the S-states are likely to be evenly distributed, so that !! should have about twice the effect of an energetically equivalent ! pH on PSII recombination a s it would affect approximately twice as many PSII centres. 94 The rate of recombination will also be affected by the redox state of the PSII electron acceptor Q A, which is also influenced by the pmf . The reduced form, Q A-, accumulates when the rate of its ph otoreduction exceeds that of re -oxidation by downstream electron carriers, feeding electrons into the recombination pathways. Activation of q E upon lowering the lumen pH as ! pH builds up, results in decreases in the PSII excitation rate and thus the fracti on of centres with Q A-. Antagonistically, pH -mediated downregulation of cytochrome b6f turnover will slow the oxidation of PQH 2, increasing the fraction of centres with reduced Q A- as electron acceptors become limited. 3.6 Control of the extent and kinetic s of pmf partitioning. At the first level, the partitioning of pmf into !! and ! pH is controlled by the biophysical properties of the chloroplast compartments. The electrical capacitance of the thylakoid membrane is small (approx. 0.6 . F cm -2) (20) so that trans -thylakoid transfer of a single electron for each PSII centre can generate a rather large !! of about 30 mV, which is equivalent to an electric field across the membrane of about 50 000 V cm -1 (152) as seen in the simulations of pmf after a s ingle turnover excitation of PSI and PSII (see Appendix, supplemental figure 3. 1a). On the other hand, the proton buffering capacity ( % ) of the lumenal compartment is quite high ( % ! 0.03 M per pH) (22), so that the same single turnover flash should produc e a very small change in lumen pH (approx. 0.001 units; Appendix , supplemental figure 3.1a). Changing the pH from 7 to 6 would require the deposition of 0.03 M of protons into the lumen (compared with approx. 10-6 M in the absence of lumen buffering groups ). These basic biophysical properties explain, in large part, why in early times (typically tens of seconds (2, 36)) after illumination, pmf is predominantly composed of !!. As illustrated in figure 3.2a, and in the simulations in figure 3.3a.1 Ða.5, in a simple thylakoid membrane with 95 no counter -ion channels, pmf remains predominantly as !! indefinitely, because !! by itself is sufficient to force protons deposited in the lumen back out through the ATP synthase, resulting in only small net changes in lumen pH. This situation is the predominant mode of action for mitochondrial and plasma membrane ATP synthase activity (reviewed in 40) and likely allows pH-sensitive biochemical processes occurring in the internal spaces to proceed unhampered by large changes in proton concentrations. As shown in the simulation, the continuous presence of high !! (figure 3.3a.1) leads to destabilization of the PSII charge pairs, while the lack of ! pH (figure 3.3a.2) prevents the formation of q E (figure 3.3a.3), resulting in accumulation of reduced QA- (figure 3.3a.4). The combination of these factors favours PSII back -reactions leading to P+680Pheo - recombination, producing large amounts of 1O2 (figures 3.3a.4 and a.5) Figure 3.2 : Illustration of the factors contributing to the balancing of pmf into !! and !pH during different phases of illumination. The blue arrows on the upper thylakoid membrane show the directions of transmembrane electron flow that generates !!, while the red arrows show the uptake and deposition of proto ns that generates ! pH. The semi -transparent arrows passing over the ATP synthase indicate the relative contributions of !! (blue) and ! pH (red) to the ATP synthase reaction. ( a) At early times after illumination, pmf is stored predominantly in !!, owing to the low electrical capacitance of the thylakoid membrane and the large proton buffering capacity of the lumen. The high !! effectively drives the efflux of protons from the lumen through the ATP synthase, maintaining a low ! pH. ( b) Activating counter -ion fluxes dissipates a fraction of !!, allowing additional proton influx and less efflux, which gradually protonates lumenal buffering groups, forming a ! pH. ( c) Counter -ion fluxes establish ion gradients that eventually reach a local equilibrium with !!, lea ding to a steady -state ratio of !!: ! pH. The generation of ! pH is then capable of downregulating cytochrome b6f turnover (triangle) as well as activating qE (dark blue circle). 96 As illustrated in figure 3.2b, adding an ion channel to the thylakoid membrane allows counter -ions to move across the membrane, down the !! gradient. For example, the simulation in figure 3.3b.1Ðb.5 shows the movement of K + (figure 3.3b.3) in response to !!, from the lumen to the stroma, progressively dissipating !! (figure 3.3b.1), while depleting the lumen of K+. Anions, such as Cl -, will also dissipate !!, but in this case, they tend to accumulate in the lumen (153). In either case, the resulting loss of !! slows the efflux of protons through the ATP synthase, allowing additional protons to be transferred to the lumen. Over time, the accumulation of protons overcomes the lumen buffering capacity, increasing ! pH (figure 3.3b.1Ðb.2) at the expense of !! (see d iscussion in 2, 76). As illustrated in figure 3.2c and simulated in figure 3.3b.1, ion fluxes cannot completely dissipate !! because the resulting counter -ion gradient will eventually prevent further movements, and the extent to which !! is dissipated will thus depend, in part, on the starting concentrations of these ions in the stroma and lumen as well as the presence of other ion transporters and proton buffers (see discussion in 2, 146). The simulation in figure 3.3b.1Ðb.5 began with stromal and lumenal K+ concentration of 40 mM, and resulted in !!: ! pH of about 1:2 in the light. It is also clear from these simulations that, even in this simplified situation, !!: ! pH can change with conditions, as is evident by the fact that !!: ! pH increases as a result of the build -up of counter -ion gradients at higher pmf (see Appendix, supplemental figure 3.1). The simulations in column C will be discussed below. In any case, allowing counter -ion flow results in dissipation of !! (figure 3.3b.1), the build -up of ! pH that acidifies the lumen (figure 3.3b.2) and activates q E (figure 3.3b.3), which decreases the fraction of reduced Q A (figure 3.3b.4). The combined effect of these changes is a decrease in 1O2 production when compared to the case with no counter -ion fluxes (figures 3.3b.4 and b.5), but at the expense of linear electron flow (LEF) (figure 3.3b.5). 97 From the above, we can conclude that the basic properties of the thylakoids allow !! to appear ver y rapidly upon illumination, whereas ! pH is formed much more slowly, with a half time on the time -scale of several minutes (2), and that these kinetics are likely to affect the onset of photoprotective mechanisms. This kinetic mismatch can be exacerbated b y the fact that at sub -saturating light, PSI and PSII centres will have access to relatively large pools of electron acceptors and donors, so that an abrupt increase in light may induce multiple turnovers, producing the large !! spikes that result in damag ing back -reactions and recombination reactions. 98 Figure 3.3 : Simulated responses of thylakoid pmf components, linear electron flow, ion fluxes and 1O2 production during a 5 -min light pulse. The simulations were performed using the DeltaPsi.py programme and initial values described in the Appendix . The timing and amplitude (maximum of 300 . mol photons m -2 s-1) of the excitation light are indicated by the light red coloured blocks. Simulations were repeated with the thylakoid permeability to counter -ions set at 0 ( a.1Ða.5), Ônormal' to approximately simulate the kinetics seen in leaves ( b.1Ðb.5), and Ôfast' (10 -fold faster than normal, c.1Ðc.5). (1) Light -induced changes in pmf (green dashed cur ves), !! (blue solid curves) and ! pH (red dashed curves), all expressed in units of volts, so that a ! pH of one is equivalent to 0.06 V. (2) The lumen pH (red, solid curves) and the relative rate constant for oxidation of PQH 2 at the cytochrome b6f complex (blue dashed curves). (3) The responses of qE (NPQ, green dashed 99 Figure 3.3 ( continued ): curves) together with the concentration of counter -ions in the lumen, [K+] (black solid curves). (4) The fraction of Q A in its reduced form (Q A-, green dashed curves) and the rate of 1O2 production (red solid curves) due to FRIP (s -1 PSII -1). (5) The cumulative LEF (green dashed curves) and 1O2 production (solid red curves). Note that the pmf parameters are shown as light -induced changes, relative to dark values. A version of this figure without this offset, supplemental figure 3.2, shows that pmf in the dark is preferentially stored in ! pH, but as pmf increases, it progressively favours !!. LEF, linear electron flow. Thylakoid properties also control the rate of ! pH relaxation (and thus q E recovery) when light is decreased, though the mechanism is more complex. As described in Cruz et al. (2, 61), when the light is switched off, electron flo w in the reaction centres is inhibited, but proton efflux through the ATP synthase continues, which causes rapid changes in !! (in the opposite direction to that induced by light -driven electron flow). Because of the low thylakoid capacitance and high lume n % , the changes in !! are far larger than ! pH, and continue until an Ôinverted' !! is established. At this point, the total pmf (!! + ! pH) is approximately equal to the backpressure from ATP hydrolysis (i.e. pmf / ! GATP/n), so the driving force for proton efflux is near zero, slowing down further proton efflux until counter -ion fluxes occur. When measuring !! changes in leaves using the electrochromic shift (ECS), this behaviour appears as the Ônegative' ECS inv phase, which is used to estimate light -induce d ! pH (2, 61), but it is important to note that these phases occur over the background level of !! from equilibration with ATP hydrolysis ( ! GATP ) in the dark, as can be seen when the simulations are plotted without offsets in the pmf parameters (Appendix , supplemental figure 3.3). Under natural field conditions in a plant canopy or an aquatic environment, light can fluctuate over a wide range of time -scales, from less than a second for wind -induced leaf movements or sunflecks and water focusing, seconds Ðminutes for changes in cloud cover, and hours for the position of the sun (66). Different regulatory processes contribute to photoprotection over these time -scales. Some photoprotection processes respond over the scale 100 of many minutes to hours (154), including the xanthophyll cycle reactions, the PSII photoinhibition/repair cycle and chloroplast movements. These slow processes are unable to respond to the more rapid fluctuations, but will approach steady -states reflecting the conditions averaged over the many minutes -to-hours time -scale. Antenna state -transitions respond over the medium times -scales, but appear to be more important in green algae than higher plants (155). Intriguingly, the remaining, rapidly responding photoprotective processes are al l controlled (directly or indirectly) by lumen pH, most importantly the adjustment of q E and the photosynthetic control of electron flow at the cytochrome b6f complex, and are thus likely to be limited by the slow rates of ! pH formation and decay. This co nclusion is in line with simulations in figure 3.3c.1 Ðc.5, which shows that increasing the permeability of the thylakoid to counter -ions by 10 -fold compared with Ônormal', increased the rate of relaxation of !! (figure 3.2c.1), leading to faster onset of !pH (figure 3.3c.1 and c.2) and thus q E (figure 3.3c.3). The overall effect is a decrease in 1O2 production (figure 3.3c.4 and c.5) owing to both a decreased !! and QA- (figure 3.3c.4). The potentially beneficial effects of increasing the permeability of th e thylakoid to counter -ion movements can be seen by comparing figure 3.2b.5 and c.5, showing that the early, rapid accumulation of 1O2 is strongly supressed as ! pH builds up. However, there is also a trade -off in loss of LEF, caused by increased control of PQH 2 oxidation (figure 3.3b.4 and c.4). The effects of counter -ion permeability on photosynthesis are highly dependent on the rates of fluctuation of the light, as is seen in the simulations in figure 4 that compare the effects of low frequency light changes (a 1 hour sine wave, figure 3.4a.1 Ð2 and b.1 Ð2) or high frequency light changes (a 1 h duration of a sine wave with period of 10 min, figure 3.4c.1 Ð2 and d1 Ð2). Overall, the lower frequency changes produced lower rates of 1O2 production and increased 101 linear electron flow (LEF) relative to the higher frequency changes, consistent with the expected frequency dependence of saturation e ffects. Increasing counter -ion permeability 10 -fold (figure 3.4b.1Ð2) had almost no effect on the simulated photosynthetic parameters under the slowly changing sinusoidal light. By contrast, the same change in counter -ion permeability had large effects und er the higher frequency square wave illumination (figure 3.4d.1Ð2), causing a twofold decrease in 1O2 under the fluctuating light, mainly due to the suppression of large !! spikes that occurred during the rapid increases in light intensity. This protection from 1O2 production, however, comes at the cost of approximately 20% decrease in LEF compared with the low frequency changes. 102 Figure 3.4 : Simulated responses of thylakoid pmf components, linear electron flow (LEF) and 1O2 production during a 1 -h light sine wave. Simulations and annotations were as in figure 3.3, but with illumination set to a 1 -h sine wave (a.1, a.2, b.1, b.2) or a 1 -h square wave ( c.1, c.2, d.1, d.2) both with peak intensity of 300 . mol photons m -2 s-1. Simulations were repeated with the thylakoid permeability to counter -ions set to Ônormal' ( a.1, a.2, c.1, c.2) or Ôfast' (10 -fold faster than normal, b.1, b.2, d.1, d.2). (a) Light -induced changes (with respect to the dark values) in pmf , !! and ! pH, all expressed in units of volts, so that a ! pH of one is equivalent to 0.06 V. ( b) The cumulative LEF and 1O2 productions. Full datasets for these simulations are presented in the Jupyter notebook in the Appendix , supplemental figure 3.3. 3.7 Can we improve photosynthesis by modifying pmf partitioning to make photosynthesis more robust? The results from our model lead us to predict that increasing the rates of counter -ion fluxes across the thylakoid could, in principle, lead to improved photosynthetic performance, 103 though the improvement i s predicted to be the long -term advantage of decreased photodamage rather than the short -term gain from increased LEF (65). Given that the slow rates of ion fluxes are likely due at least in part to low protein levels of ion transporters in the thylakoids , a strategy of overexpressing the rate -limiting channels could be suggested. However, if better photosynthesis could be achieved this simply, plants might be expected to have already evolved more rapid ion fluxes than those measured in laboratories. Indee d, exploring these properties in natural populations may reveal precisely these sorts of variations. On the other hand, increasing ion fluxes may lead to secondary, deleterious effects. A basic tenet of the model is that ! pH cannot form without the moveme nts of counter -ions, and thus, for each proton that accumulates in the lumen, approximately an equal number of counter -ion charges must be moved to dissipate the !!. In effect, energy stored in !! is consumed, at least temporarily, in the movement of count er-ions, removing the ATP synthase driving force and preventing proton efflux through the ATP synthase. The diversion of protons away from ATP synthase efflux then allows protons to enter the lumen buffering pool, allowing ! pH to form more rapidly as proto ns are no longer being driven out of the lumen by !! and the buffering capacity is overcome. Thus, one consequence of ! pH formation will be a transient decrease in the LEF output ratio of ATP/NADPH due to the time -dependence of overcoming the lumen bufferi ng capacity. Metabolic congestion and photodamage can occur if this ratio does not precisely match that needed to power assimilation, requiring alternative electron transfer processes, such as cyclic electron flow or the water Ðwater cycle, to make up the b alance (6). Our simulations suggest that a slow ! pH formation (half time of about 4 min) results in a counter -ion related deficit of about 0.05 ATP per CO 2 fixed by assimilation, which should be easily remedied 104 by alternative electron transfer processes (f igure 3.5). With a 10 -fold faster ! pH formation provided by increased counter -ion flux, the resulting ATP deficit caused by counter -ion movements can become severe, requiring a 10 -fold higher input of ATP per CO 2 fixed (figure 3.5c), which may exceed the c apacity for cyclic electron flow in some species. Figure 3.5 : Effects of thylakoid counter -ion fluxes on the ATP/NADPH budget of photosynthesis. Data are taken from the simulations performed in figure 3.3, with x-axis origin set to the beginning of ill umination. ( a) The light -induced flux of counter -ions (K +), with positive values representing net flux out of the lumen. ( b) The proton deficit, i.e. the protons deposited into the lumen but buffered so that they are unavailable to the ATP synthase. ( c) The cumulative deficit in molecules of ATP relative to PSI turnover needed for CO 2 fixation by the Calvin ÐBenson cycle. The red, yellow, green and blue lines represent results with the thylakoid counter -ion permeability set to zero, Ônormal' (figure 3.3), 10) normal and 100 ) normal, respectively. The osmotic balance of the chloroplast compartments must also be finely tuned to maintain their structure and the function of proteins residing within each compartment, and even small osmotic imbalances can lead to swelling -induced loss of thylakoid stacking, or shrinkage -induced inhibition of the interactions of plastocyanin with PSI (2, 153, 156). Proton translocation 105 by itself should not appreciably affect the osmotic potential of the lumen because most protons are buffered. By contrast, counter -ion movements will very likely change the concentrations of free counter -ions, and thus have a colligative effect on the osmolarities of the lumen and stroma. It is interesting to note that loss of the chloroplast potass ium KEA (the potassium Ðproton antiporter in the thylakoid membrane) transporters leads to swelling and disordering of the thylakoid structure (29), suggesting that fine -tuning of the thylakoid ion balance is critical for osmoregulation. It is also suggesti ve that Chlamydomonas cells are able to compensate for severe hyperosmotic shock -induced lumenal shrinkage, but only over the same time -scale as ! pH formation, i.e. about 5 Ð10 min (157). It is thus possible that the rate of pmf partitioning is limited by t he need to prevent acute osmotic imbalances that could result in structural perturbations. At this point, the model is not intended to reproduce all the reactions of photosynthesis. Nevertheless, using reasonable, published values for thylakoid properties (see Appendix ), the model qualitatively reproduced the FRIP behaviours observed by Davis et al. (36), including light fluctuation -induced !! spikes, that result in increased recombination, 1O2 production and the accumulation of photodamage. This simplifi ed model also suggests that accelerations in counter -ion fluxes will result in tuned increases in the rate of NPQ onset, which will have a potentially beneficial effect by decreasing 1O2-related photodamage and accelerating q E responses, but this will be a t the cost of decreasing LEF. This suggestion is interesting in the light of the recent work of Kromdijk et al. (126), who reported that more rapid NPQ responses can increase plant yield by increasing PSII quantum efficiency (and thus LEF). Our simulations suggest an alternative explanation: that the underlying benefit of more rapid responses will be decreases in both ROS production and 106 photodamage, leading to more sustained photosynthesis and lower input costs over the long term. Interestingly, further mod ifications of photosynthetic parameters in our simulations show that the trade -off loss of LEF can be avoided by decreasing the p Ka for controlling the cytochrome b6f complex to well below that for initiation of q E, but this may lead to less control of ele ctron flow at the cytochrome b6f complex and over -reduction of PSI and subsequent PSI photodamage (5). At the very least, recent results and the simulations they inspire suggest possible targets for plant improvement, and generate testable hypotheses. Ult imately, improving photosynthesis will require understanding the multiple constraints that life in the real world imposes on photosynthesis, as well as the multiple, interacting regulatory systems that have evolved to cope with them. Approaching this compl ex problem will require a larger scale investigation of the responses of pmf in a range of species, and under field -like conditions, as well as a deeper understanding of how the biophysical machinery of photosynthesis is integrated with the host organism t o respond to the challenges of rapidly fluctuating environmental conditions. From the simulations presented above, taking steps towards such an integrated view can reveal emergent properties of the system that were not apparent from studies of isolated complexes. Towards that end, it is hoped that future work (by us and others) will expand the model presented here to test the effects of important processes, including newly discovered ion transport systems and their regulation (158, 159), effects on thylakoid osmotic balance (160), the PSII damage/repair cycle (161, 162), the accumulation of electrons on the acceptor side of PSI (which can lead to irreversible PSI photodamage (163)), alternative modes of PSII regulation including the rece nt report of bicarbonate -mediated protective redox tuning (132), regulation of the ATP synthase, and the need for alternative electron transfer pathways. 107 Holistically, an expansive mechanistic model of photosynthetic regulation will likely provide immedia te targets for testing the effects of engineered changes aimed at improving photosynthetic yields. When the model fails to replicate experimental results, it has the potential not only to focus attention on gaps in our understanding of photosynthesis and i ts regulation but also to provide insights that could help to fill these gaps. 3.8 Acknowledgements. Work performed by G.A.D. and D.M.K. was supported by the US Department of Energy (DOE), Office of Science, Basic Energy Sciences (BES) under award number DE-FG02 -91ER20021. A.W.R. was supported by the Biotechnology and Biological Sciences Research Council grant nos. BB/K002627/1 and BB/L011206/1 and the Royal Society Wolfson Research Merit Award. 108 Supplemental Figure 3.1: Simulations of the amplitudes of pmf parameters induced by a single -turnover flash hitting all photosystem II (PSII), but no photosystem I (PSI). The simulations were set up to recapitulate the experiment performed in Davis et al. (36) (c.f. Figure 6). To obtain single turnover condition s, the rate constant for re -oxidation of Q A- was set to zero. Excitation of PSI was inhibited by setting its antenna size to zero. In Panels A.1 and A.2, the fluxes of counterions were also set to zero to reveal the electrogenic effects of both excitation and recombination. In Panels B.1 and B.2, the permeability of the thylakoid to counterions was set very high to simulate the effects of addition of KCl + valinomycin to rapidly dissipate flash - induced !!. Panel A.1 and B.1 show the effects on !! (blue sol id line), !pH (red dashed line) and total pmf (green solid line). Panels A.2 and B.2 show the predicted concentration of the S2QA- (green dashed line) and 1O2 production (red dashed line). Panels A.3 and B.3 show the cumulative LEF and 1O2 production. As s een in Davis et al. and references within (36), the presence of a !! induced more rapid 1O2 production. However, because all charge -separated states recombine through the same pathway, the cumulative levels of 1O2 production for both conditions were the same. 109 Supplemental Figure 3.2: Simulated responses of thylakoid pmf components, linear electron flow, ion fluxes, and 1O2 production during a 5 -min light pulse. The simulations are as in Figure 3.3, but the pmf parameters are not offset at time zero. 110 Supplemental Figure 3.3: Simulated responses of thylakoid pmf components, linear electron flow, and 1O2 production during illumination with a 1 -hour sine or square waves. The figure contains the full d ata set for the simulations described in Figure 3.4 in the main text. Simulations were performed as in Figure 3.3, but with illumination set to a one -hour sine wave (Panels in columns 1 and 2, starting with A, B) or a one -hour square wave (Panels in column s 3,4, starting with C, D) both with peak intensity of 300 µmol photons m -2s-1. Simulations were repeated with the thylakoid permeability to counter -ions set to ÒnormalÓ (Panels in columns 1, 3, starting with A, C) or ÒfastÓ (10 -fold faster than normal, pa nels in columns 2, 4, starting with B, D). The panels in row 1 show light -induced changes (with respect to the dark values) in pmf , !!, and !pH, all expressed in units of V, so that a !pH of one is equivalent of 0.06V. Row 2 shows the effect of the lumen pH on the turnover rate of cytochrome b6f. Row 3 shows the responses of qE together with the concentration of counter -ions in the lumen, [K +]. Row 4 shows the fraction of Q A in its reduc ed form (Q A-) and the rate of 1O2 production due to FRIP (s -1 PSII -1). Row 5 shows the cumulative LEF and 1O2 productions. 111 Chapter 4 The electric field component of the photosynthetic proton motive force increases the yield of PSII recombination in vivo during light fluctuations Geoffry A. Davis and David M. Kramer 112 4.1 Abstract Photosystem II (PSII) functions as a water -plastoquinone oxidoreductase to mediate the capture and conversion of light into chemical energy during photosynthesis. This reaction mechanism requires two successive PSII turnovers to reduce plastoquinone and four turnovers to oxidize water, requiring coordination of multiple electron transfer steps occurring on the donor and acceptor sides of the protein complex to occu r cooperatively for proper functioning. However, reversal of electron transfer (recombination) can occur, leading to side -reactions that produce a high yield of triplet chlorophyll and singlet oxygen, which result in irreversible oxidative damage and inact ivation of PSII. These recombination events are suggested to be modulated in vivo by the extent of the electric field ( !!) across the thylakoid membrane, which destabilizes charge -separated states within PSII proportionally to the vectorial distance across the membrane. To identify the impact of !! on PSII recombination in vivo , delayed fluorescence was measured in intact Arabidopsis thaliana during fluctuations in light intensity to induce rapid, transient large !!. Rapid increases in light intensity induced large, transient increases in the yield of PSII recombination, while static, high light did not increase PSII recombin ation yields. Using mutants deficient in activating photoprotective quenching mechanisms, PSII recombination occurs over a longer time than in wild type plants. Taken together, these results suggest that fluctuating light rapidly increases the yield of PSI I recombination, which then decays relative to the depletion of !! and the concentration of PSII species available to recombine. The increases in PSII recombination under fluctuating light may explain the susceptibility of photosynthesis in natural, field conditions where rapid light fluctuations occur throughout the course of the day. 113 4.2 Introduction Oxygenic photosynthesis in cyanobacteria, algae, and plants utilizes electrons generated from the oxidation of water to ultimately yield NADPH and ATP for cellular anabolism. This is accomplished by harvesting light in antenna pigments and funneling the excitation energy to special pair chlorophylls (P) within each of photosystem II (PSII) and photosystem I (PSI). PSII is a water Ðquinone oxidoreductase, utilizing the energy generated via excitation reaction center chlorophyll to oxidize water and reduce plastoquinol (PQH 2), releasing the protons into the thylakoid lumen during the water splitting process (164). The oxidation of PQH 2 by the cytochrome b6f complex leads to additional proton deposition into the thylakoid lumen through the Q -cycle (10) and electron transfer to plastocyanin, which is used to re -reduce PSI following excitation and charge separation (165). Electrons from PSI are terminally transf erred to ferredoxin and subsequently to NADP + to generate NADPH (127). The deposition of protons into the lumen during electron transfer events generates a light -induced ! pH across the thylakoid membrane. Acidification of the thylakoid lumen triggers regu latory processes that limit electron transfer at PSII. The availability of oxidized plastoquinone (PQ) for PSII electron transfer is limited by the rate of PQH 2 oxidation by the cytochrome b6f complex, the turnover of which slows by a factor of 10 per pH u nit decrease (11, 12), increasing the reduction state of the PQ pool and therefore limiting quinone substrate availability for PSII. This is counterbalanced in algae and higher plants by pH -mediated activation of the energy -dependent (q E) form of nonphotoc hemical quenching (NPQ), which consists of protonation of the thylakoid membrane protein PsbS (7), as well as violaxanthin de -epoxidase protonation in the thylakoid lumen to activate the conversion of violaxanthin to zeaxanthin (166). Both components of q E ultimately lead to excitation energy quenching in the 114 antenna pigments and decreased excitation pressure on PSII (6). Dissipation of ! pH occurs predominantly via proton translocation by the thylakoid ATP synthase to produce ATP (40), however the critical activities of ion channels within the thylakoid membrane to modulate the partitioning of the transthylakoid proton motive force ( pmf ) between a ! pH and electric field (!!) are increasingly being identified (34, 35, 167). The multifaceted complexity of electron transfer and pmf reactions across the thylakoid membrane in vivo leads to particular stresses on PSII. PSII is a multi -protein complex where the redox chemistry occurs across the membrane Ðbound reaction center core D 1 and D2 subunits. Following the transfer of excitation energy from antenna pigments to PSII, excitation of chlorophyll molecules within the PSII (P*) core leads to primary charge separation through electron transfer from P 680 to a nearby pheophytin (Pheo) molecule to form the P 680+Pheo - state (Fig. 4.1). The initial charge separation is stabilized by subsequent electron transfer to the permanently bound quinone Q A, and then to a secondary quinone Q B. Re-reduction of P 680+ is mediated by a redox active tyro sine (Y Z) that participates in the transfer of electrons from the PSII lumen -exposed oxygen -evolving complex (OEC), which goes through a series of at least four redox intermediates (S -states) to produce O 2 (17, 164). PSII functions as a two -electron gate, requiring two successive PSII turnovers to fully reduce the quinone bound at Q B, whereupon the quinol diffuses out of the PSII Q B site into the thylakoid membrane. This sequential reduction of Q B requires the semiquinone, Q B-, formed after the first PSII turnover to be held stably through a second PSII cycle. The forward ÔproductiveÕ electron transfer routes are thermodynamically favorable, however, the reversal of charge Ðseparated states, termed recombination, can occur due to various factors, and are mediated by the free -energy gap between the corresponding states, the physical distance between 115 redox carriers, and the reorganizational energy (56, 168). While electron back -reactions limit productivity by decreasing the overall efficiency of photosynthesis , PSII back -reactions are precarious in vivo due to the recombination of P 680+QA- through Pheo as an intermediate, described as the indirect recombination pathway due rather than the direct recombination of Q A- to P 680+. Indirect recombination from Q A-, or QB- back to Q A- due to the relatively small equilibrium constant for sharing electrons between Q A and Q B (88), can generate both the initial charge separated singlet state ( 1[P680+Pheo -]) as well as produce a high yield of the triplet state (3[P680+Pheo -]) (141). The 3[P680+Pheo -] state decays to the triplet ground state chlorophyll ( 3P), which has enough energy to interact with O 2 to generate singlet oxygen ( 1O2), which is highly reactive and proposed to irreversibly damage the D1 protein and lead to irr eversible PSII photoinhibition (67, 96), requiring degradation of the damaged D1 and de novo synthesis of a new D1 protein to repair the damaged reaction center (128). 116 Figure 4.1: Photosystem II electron transfer reactions successively increase the dist ance of transmembrane charge separation. Electron transfer within the PSII reaction center core D1 and D2 subunits occurs vectorially across the span of the thylakoid membrane (A). Light is capture in pigments within antenna proteins and the excitation ene rgy funneled to the reaction center. Excitation of the core PSII chlorophylls (P *) leads to the initial charge separated state P 680+Pheo - along the D1 branch. The electron is subsequently transferred from Pheo D1 to the bound quinone Q A, followed by electro n transfer to the terminal PSII electron acceptor Q B. Re-reduction of P 680+ is accomplished via the extraction of electrons from the Mn 4CaO 5 oxygen evolving complex (OEC) S -states through a redox active tyrosine (Y Z). Secondary electron transfer reactions thermodynamically stabilize the productivity of forward electron transfer through successive electron transfer steps to lower potential redox carriers (B). Reversibility of electron transfer reactions (B, dashed lines) are mediated by the free energy diffe rence between the charge separated species. Increasing transmembrane !! destabilizes the vectorial movement of charges across the membrane relative to the distance between cation and anion species. For simplicity this is illustrated for the effect on P+QA- but will result in changes proportional to the distance between membrane bound charge separated species. A second PSII turnover is required to reduce Q B- to Q BH2, where it can then diffuse out of the Q B site and exchange for oxidize PQ. Positions of PSII cofactors based on (169). + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + +++++- - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - !! Pheo D1 QA QB ChlD1 PD1 PD2 OEC YZ Pheo D2 A B !! !G !G P* P 3P 1[P+Pheo -] 3[P+Pheo -] P+QA- P+QAQB- Sn+1QAQB- Sn+1QA- P+QA- Standard Free Energy (V) 117 Reaction center back -reactions have been well studied in vitro , where the free energy gap between states can be experimentally manipulated. In isolated reaction centers or isol ated thylakoid membranes, increases in the rate of electron recombination occurs when the free energy gap between P +QA- and P +Pheo - (where P is the primary donor) is altered by substituting exogenous quinones (142), utilization of a salt -induced !! (26, 170), or application of external electric fields (171, reviewed in 102). In vivo , !! has been correlated with an increase in 1O2, PSII damage, and photoinhibition (36), suggesting that in vivo the free energy gap of charge separated states in PSII are modula ted by the partitioning of pmf , and the rate of electron recombination increases in vivo due to destabilization of charge separated states via !! (38, 172). To directly test this possibility, millisecond timescale delayed fluorescence has been utilized to monitor PSII back -reactions in vivo . Delayed fluorescence (also referred to as delayed light emission) originates from repopulation of the excited P * via recombination reactions from electrons already trapped by PSII (173, 174). Repopulation of P 680* requ ires back -reaction from 1[P680+Pheo -], which can be generated by recombination from P 680+QA-. However, as discussed above, recombination from P 680+QA- to P 680+Pheo - generates both the singlet and triplet states, the latter of which forms with a higher yield (141). Therefore, monitoring the delayed fluorescence emission provides discreet information on the rate of PSII charge recombination as well as an estimate of th e production of 3[P+Pheo -] during these back -reactions. Along with the decreased frequency of forming 1[P680+Pheo -] via P 680+QA- recombination relative to the triplet state, 1[P680+Pheo -] can directly decay to the ground state non -radiatively rather than t hrough the energetically uphill repopulation of P *. Although the emission of light as delayed fluorescence is approximately 100 -fold lower than that of prompt fluorescence (175), its utility as a probe of 118 PSII recombination reactions in vivo . Utilizing del ayed fluorescence as a probe of PSII recombination in vivo under environmental conditions can provide insight into mechanisms influencing PSII electron transfer reactions and photodamage in vivo , enhancing the capacity to predict stressors that will alter PSII photodamage rates through changes in electron recombination reactions. 4.3 Materials and Methods 4.3.1 Plant and growth conditions. Arabidopsis thaliana plants were grown on soil at 21¡C with 100 µmol photons m -2s-1 light under a 16:8 day:night cycle . Spectroscopic measurements were performed on plants three weeks after germination. Arabidopsis thaliana mutants lacking PsbS ( npq4) (176) and vio laxanthin de -epoxidase ( npq1) (166) are both in the Col -0 background. 4.3.2 Spectroscopic measurements of de layed fluorescence, prompt fluorescence, and the electrochromic shift. Measurements of chlorophyll fluorescence and the electrochromic shift were performed using a custom made spectrophotometer described previously (117) with modifications to utilize three detectors during each measurement. Individual detectors were used to measure electrochromic shift (ECS) transmission, the reference intensity of the measuring pulse prior to leaf absorption, and a photomultiplier tube ( PMT) to measure both prompt and delayed fluorescence. Red actinic illumination was used in all experiments to minimize the effects of chloroplast movement on chlorophyll fluorescence parameters, as red light does not induce chloroplast movements (119). All measurements were performed under at room temperature under ambient conditions. 119 The ECS was measured at 520 nm using detectors positioned in line with the actinic and measuring light, and collected after passing through a BG18 bandpass filter as describ ed previously (74). The amplitude and decay kinetics of the ECS were determined by fitting the dark interval relaxation kinetics to a first -order exponential decay to determine the total proton motive force (ECS t) and the conductivity of the ATP synthase t o protons ( gH+) (74). The ECS was corrected for pigment variations by normalizing to the total chlorophyll per leaf area (36). Estimates of the redox state of Q A (qL) was determined 5 s, 15 s, and 30 s following each light transition (100). Delayed chlorop hyll fluorescence was measured using a PMT ( R636-10, Hamamatsu, Inc.) with a commercially available socket assembly (C1392 -57, Hamamatsu, Inc.) . Delayed fluorescence was measured by interrupting the actinic illumination every 5 s with a 130 ms dark interva l. During the 130 ms interval, data was sampled from the PMT every 200 us. The prompt fluorescence decay and ECS during the dark interval were measured simultaneously every 1ms with a 10 µs 520 nm measuring pulse. The timing and intensity of the measuring pulse were found to not alter the yield or kinetics of the delayed fluorescence signal. 4.3.3 Determination of delayed fluorescence yields. Both delayed and prompt fluorescence originate from the same antenna pigments (177, 178), and therefore the relativ e yields of prompt and delayed fluorescence at a given time are equally impacted by the quenching processes occurring in the antenna at the time, e.g. the probability of measuring a photon emitted as delayed fluorescence is equal to the probability of measuring a photon emitted as prompt fluorescence under the same quenching conditions with the same detection unit. Therefore, the delayed fluorescence of each dark interval was normalized to the prompt fluorescence during the same dark interval by interpolati ng the prompt 120 fluorescence decay to each of the delayed fluorescence measurements in order to determine the yield of delayed fluorescence. The normalized delayed fluorescence signal was then integrated over the dark period to determine the delayed fluoresc ence yield. At each light intensity, a baseline offset was found to have occurred in the PMT proportional to the light intensity. Therefore the average of the last 5 measurements at each light intensity were subtracted from the calculated delayed fluoresce nce yields to proportionally correct each measurement for the offset caused by the actinic illumination. 4.4 Results 4.4.1 Abrupt increases in light intensity increase the yield of delayed fluorescence. To determine the effect of light fluctuations on the delayed fluorescence yield ( #DF), plants were subjected to a series of fluctuating actinic intensities, during which delayed fluorescence, prompt fluorescence, and ECS parameters were measured nearly sim ultaneously throughout the illumination period (Fig. 4.2). If P 680* is repopulated via PSII recombination in the dark, relaxation to the ground state is accompanied by the release of the exciton back into the antenna pigment bed and can be measured as the delayed fluorescence signal if it is emitted as fluorescence by the antenna. The probability of measuring delayed fluorescence will not only depend upon the repopulation of P 680*, but also upon the probability that after equilibration with the antenna pigm ents that excitation is not quenched but instead emitted as fluorescence. Therefore #DF can be determined by normalizing the delayed fluorescence to the prompt fluorescence at the same measurement time, as the probability of the antenna emitting a photon as fluorescence will be equivalent for both prompt and delayed fluorescence at the same given time. 121 As predicted by Davis et al. (36), rapid changes in light intensity produced short, rapid increases in the yield of delayed fluorescence when the light inte nsity is increased (Fig. 4.2A), indicating transient periods where PSII recombination reactions are increased. These increases in delayed fluorescence presumably arise from large, transient increases in !! across the thylakoid due to the low electrical cap acitance of the thylakoid membrane (2, 20, 36). As can be seen from the pmf measured concurrently with the delayed fluorescence (Fig. 4.3B, D), increases in light intensity leads to a near instantaneous increase in the total pmf . While light fluctuations from one light intensity to a lower intensity were not expected to increase #DF due to the field induced recombination model, the rapid decrease in excitation pressure did not show a significant change in yield relative to the proceeding steady state yield s even though the PSII excitation is rapidly decreased. While it has been shown that photosynthesis is sensitive to light fluctuations (36, 60, 66, 75, 179), PSII photoinhibition is known to be linearly dependent upon the light intensity (78). As such, #DF was measured at 400 µmol m -2s-1 (Fig. 4.2B), approximately four time s the growth light intensity, which is the average light intensity of the fluctuating light treatment. Similarly to the fluctuating light (Fig. 4.2A), upon the initial dark -light transit ion #DF increases before decreasing to near the detection limit. However, unlike under fluctuating light conditions, even at four times the growth light intensity and the same total quanta of the fluctuating light treatment, the yield of delayed fluorescen ce under constant light does not increase after the initial dark -light transition, and remains near the detection limit throughout the illumination period. 122 Figure 4.2: Fluctuating light increases PSII recombination in vivo . Delayed and prompt fluoresc ence were measured every 5 s durin g 100 ms dark intervals in wild type Col -0 Arabidopsis thaliana leaves. For each dark interval, the yield of delayed fluorescence (#DF) was determined by integrating the DF/PF signals at each time point. During fluctuating light (A), transient increases in #DF occur immediately after each low light/high light transition. Following dark adaptation, light intensities were maintained for 14 minutes at 100, 600, 200, 1000, and 100 µmol m -2s-1 (yellow bars). Under static 400 µmo l m -2s-1 light (B), #DF increases only during the initial induction following dark adaptation (mean + s.d., n=4). 4.4.2 Alterations in the regulation of PSII Q A redox state leads to changes in field -induced increases in recombination. The redox state of Q A, determining the availability of substrate Q A- for PSII recombination, is decreased in vivo by the regulation of exciton transfer from the antenna pigments to PSII cores by NPQ processes. In the Arabidopsis npq4 mutant, which lacks PsbS, the pH -dependent q E component of NPQ is severely compromised (176), leading to an increase 123 in the excitation pressure on PSII under conditions that would normally be limited by q E. When subjected to fluctuating light (Fig. 4.3A), npq4 plants experience increased PSII recombination relative to Col -0, reflected by a statistically higher #DF. Unlike Col -0, where the electric field induced increases in recombination decrease to background level in approximately 1 -1.5 minutes, the increase in delayed fluorescence yiel d in npq4 persists for 2.5 -7 minutes. The long -lived increase in #DF does not appear to be a factor of an increase in light over growth intensity, as at 200 µmol m -2s-1 (2X growth light), there are not difference s between npq4 and Col -0, both of with have #DF near the detection limit. Arabidopsis plants lacking violaxanthin de -epoxidase (! VDE, npq1) are also deficient in q E quenching, though not to the extent of npq4 (180). Similarly to npq4, npq1 plants exhibit ed a prolonged increase in #DF upon increases in light intensity, though the increased yield decreases faster than in npq4 (Fig. 4.3C). Except for the transition from a dark adapt ed state, where the pmf stays elevated relative to Col -0, the total pmf between Col -0 and the two q E mutants was not statis tically significantly different following the light fluctuations (Fig. 4.3B, D), highlighting that the diminished ability to regulate excitation energy transfer to PSII to limit the accumulation of Q A- increases the recombination frequency relative to wild type. 124 Figure 4.3: Deficiencies in pH -dependent NPQ antenna quenching increase # DF due to increased PSII center with Q A-. Changes in PSII recombination due to an i mpaired abilit y to induce pH -dependent q E quenching in Arabidopsis npq4 (red, A and B) and npq1 (blue, C, D) w ere assessed under fluctuating light. Light fluctuations were performed as in Fig. 4.2A. Col -0 (black) #DF from Fig. 4.1 shown as reference comparison. Increases in #DF are seen in both npq4 (A) and npq1 (B) following increases in light intensity (yellow bars). Total pmf (ECS t) measured at the same time as #DF is not different from Col -0 following increases in light intensity in either npq4 (B) or npq1 (D). The fraction of centers with Q A- (qL) (E) measured 5, 15, and 30s after each ligh t fluctuation is increased in the q E mutants after transitions to high light (mean + s.d., n = 3-4) (*, p < 0.05 ). 4.5 Discussion Regulation of the light reactions of photosynthesis is critical for prolonged maintenance of photosynthetic capacity. Multi ple mechanisms have evolved to regulate PSII excitation pressure (57, 62) to limit photooxidative damage and the reduced photosynthetic capacity 125 incurred when reaction centers are inactivated (65). Even with multiple levels of regulation, photodamage to PSII leads to the rapid turnover of the reaction center D1 protein (128, 181). While multiple mechanisms have been proposed to explain the cause(s) of PSII photodamage, most center on the generatio n of reactive oxygen species by PSII side reactions and subsequent protein damage caused by ROS (56, 67, 68). In the Arabidopsis thaliana minira mutants (36), alteration of the steady -state !! lead to an increase in 1O2 production and PSII damage in vivo . This lead s to the proposal that a large !! in vivo , experienced near continuously in minira mutants as well as during light fluctuations in wild type plants, can modulate PSII back reactions in vivo and increase the frequency of Q A- recombination, increasi ng 1O2 production. To test this hypothesis, we utilized the yield of delayed fluorescence as a direct indicator of changes in PSII recombination reactions in vivo . The high (near unity) quantum efficiency of PSII under optimal conditions leads to success ful charge separation for nearly every exciton that reaches PSII (182). While the redox intermediates within PSII are poised to favor forward electron transfer reactions through successive decreases in midpoint potential and an increase in the donor/accept or distance, non -productive recombination reactions can occur. While various methods have been employed to understand the energetics of PSII chemistry through mediating recombination reactions (reviewed in 174), we have utilized delayed fluorescence emissi on to monitor PSII recombination as it allows intact plants to be measured over prolonged periods of time. This allowed near continuous measurements of delayed fluorescence during the course of light treatments on intact Arabidopsis leaves. Dark repopulati on of P 680* and emission of light as delayed fluorescence was observed in all plants tested upon a shift to higher light intensity (Figs. 4.2 and 4.3). Upon a high light 126 transition, an immediate increase in the pmf is observed (Fig. 4.3), which is predomin antly held as !! due to the low electrical capacitance of the thylakoid membrane (2, 36, 167). The rapid increases in #DF decrease within minutes to minimal levels, likely responding to the combinatorial impact of a change in pmf partitioning that occurs f rom ion transport across the membrane to dissipate !! as well as the onset of q E to decrease the fraction of PSII with Q A- (167). The generation of 1O2 via PSII acceptor side back reactions and 3P production has been shown to correlate with the rate of PSII photoinhibition (36, 183, 184). Traditionally, PSII photoinhibition has been thor oughly characterized in a light -dependent manner, with increased photoinhibition being linearly dependent upon t he light intensity (78). Similarly, 1O2 generation has been shown to linearly increase with actinic illumination (185), and the overall 1O2 production to be dependent upon the P 680+QA-/P680+Pheo - free energy gap (184, 185). Contrary to a linear light induc tion curve, the fluctuating light used above oscillates between lower and higher light intensities. As suggested by Davis et al. (36), low to high light transitions favoring transient !! spikes increased the yield of PSII recombination. However, an increas e in irradiance over growth light (200 µmol m -2s-1 vs. 100 µmol m -2s-1) did not produce differences in #DF, consistent with a lack of !! generation during these transitions. Photosynthesis is sensitive to natural light conditions, where rapid transitions in the available light intensity can occur throughout the day (66). Intriguingly, under standard laboratory (i.e. non -fluctuating) light conditions, the lack of regulating PSII excitation in npq1 and npq4 does not lead to a seed yield penalty, however grow th under variable light in either a laboratory or field decreased seed yield in both npq1 and npq4 (66). During fluctuations in actinic light, both npq1 and npq4 experienced sustained increases in #DF relative to Col -0 (Fig. 127 4.3), indicating prolonged incr eased yields of PSII recombination through the indirect, 3P generating pathway, potentially explaining the sensitivity of these mutants to long term growth under fluctuating light. The electron transfer reactions of PSII occur on the order of nanoseconds to microseconds (164), requiring coordination of reactions occurring on both the donor and acceptor sides of PSII at different rates. The acceptor side of PSII is limited by the rate of Q A-/QB electron transfer and the availability of oxidized PQ to bind a t the Q B site (186). As ! pH increases during photosynthesis, the turnover rate of cytochrome b6f decreases (11, 12), shifting the PQ/PQH 2 ratio and limiting the availability of oxidized PQ, increasing the lifetime of Q A- if the quinol exchange range becomes limiting, and increasing the likelihood of P 680+QA- recombination. Concurrently, activation of q E via PsbS and VDE protonation as the ! pH increases should relieve some of the excitation pressure on PSII by quenching ex citation energy in the antenna pigments rather than transferring it to PSII. However, the electron transfer rates of PSII are much faster than the buildup of ! pH (2), which initially requires overcoming the buffering capacity of the thylakoid membrane (22). The generation of !!, however, can be generated rapidly, as the low electrical capacitance of the thylakoid membrane leads to the generation of !! via the trans -thylakoid movement of electrons by the reaction centers (20). This kinetic mismatch in the ti ming of !! generation versus the pH -dependent processes that help protect reaction centers from over -excitation (discussed in 167) has been proposed to increase the rate of PSII photodamage and the generation of 1O2 (36). Utilizing delayed fluorescence as a direct indicator of the yield of PSII recombination, the same conditions that generate rapid increases in pmf and !! leads to enhanced PSII recombination. While the generation of !pH lags compared to !! generation following a high -light fluctuation, resu lts from mutants deficient 128 in q E induction suggest that regulating the PSII excitation pressure in steady -state to minimize the concentration of Q A- decreases #DF immediately after the light fluctuation due to the decreased availability of states available to recombine. Recombination from Q A- in PSII is dependent upon four main factors: 1) presence of Q A-, 2) the redox state of the quinone at the Q B site, 3) the free energy difference between P 680+QA- / P680+Pheo -, and 4) the PSII donor side existing in an S-state capable of recombination from Q A- (S2 and S3) (187). Under continuous illumination, due to PSII misses that fail to advance the S -state (17), within seconds the population of PSII within the leaf will become asynchronous between the S -states, with each state being approximately equally represented. This leads to approximately 50% of PSII being capable of Q A- recombination at a given time due to the donor side availability (167). The free energy difference between P 680+QA- and P 680+Pheo - is modulate d by the trans -thylakoid !!, which can be modified by the movement of ions across the membrane, but is large following abrupt changes in light intensity (2, 36). These large !! spikes are enough to increase the recombination frequency from Q A- in wild type Col-0 plants (Fig. 4.1A). Characterization of specific recombining states during near Ðcontinuous measurements of delayed fluorescence, both as performed in the current work or with a phosphori scope, has proven difficult to ascertain . Due to the total PSII population becoming asynchronous between both the PSII donor side and acceptor side reduction states, multiple pathways for Q A- recombination, as well as the continuation of forward electron transfer during the measuring interval if the Q B site is not blo cked , delayed fluorescence under continuous illumination is the result of a multitude of states (188). The rates of PSII donor side electron transfer reactions vary depending on the S -state, however, within 10 µs 80% of PSII will progress from Y ZP680+ to 129 YZ+P680 (189), and within ~1 ms reduction of Y Z+ and advancement of the S -state will have occurred (16). Therefore, after the first 1 -2 measurements of delayed fluorescence obtained in our experiments, re -population of P 680* will require equilibrium reactio ns occurring on both on the donor and acceptor sides of PSII. However, the yield of delayed fluorescence represents a fraction of the total PSII recombination routes, which primarily proceed via repopulation of P680 +Pheo - (145) (Fig 4.1). When recombining via this indirect route, van Mieghem et al. found 66-75% of Q A- recombination occurred by generation of 3[P680+Pheo -] rather than 1[P680+Pheo -] at 20K (141). Using an externally applied electric field, de Grooth and van Gorkom found that only ~3% of Q A- recombination repopulated P 680*, suggesting that the delayed fluorescence measured in the current experiments during light fluctuations represents less ~5% of the total yield of PSII recombination, the majority of which is presumed to proceed via a route tha t favors generation of 3P. Under these conditions, if the assumption that a large increase in 3P and 1O2 generation is correct (36), an increase in PSII photodamage should occur rapidly after a low light to high light transition, which could explain the in creased sensitivity of photosynthesis to natural light conditions. As delayed fluorescence can be measured non -destructively, an increased understanding of the relationship between delayed fluorescence and PSII damage will allow the targeted manipulation o f photosynthetic processes to minimize the detrimental effects of the !!-mediated increases in PSII recombination following light fluctuations while maintaining photosynthetic efficiency. 4.6 Acknowledgements Work performed by G.A.D. and D.M.K. was supported by the US Department of Energy (DOE), Office of Science, Basic Energy Sciences (BES) under award number DE -FG02 -91ER20021. 130 Chapter 5 Natural variation in the activation and dissipation of nonph otochemical quenching in Arabidopsis thaliana subjected to fluctuating light Geoffry A. Davis, John E. Froehlich, David M. Kramer 131 5.1 Abstract Improving the productivity of photosynthesis in natural, field conditions requires an improvement in our current understanding of factors mediating the onset and dissipation of photosynthetic regulatory processes. Feedback regulation of the light reactions of photosynthesis is mediated by the buildup and subsequent dissipation of a light -induced pH gradient ( ! pH) across the thylakoid membrane, which subsequently induces nonphotochemical energy quenching (NPQ) by light -harvesting antenna pigments and downreg ulates the rate of electron transfer through the photosystems. The kinetics of ! pH generation and photosynthetic repression combined with the kinetics of removing these limitations leads to periods of light availability that are not being optimally utilize d by photosynthesis for energy production during natural, fluctuating light conditions. To identify genetic factors mediating variation in the onset, dissipation, and total steady -state NPQ during light fluctuations, the Arabidopsis thaliana multi -parent a dvanced generation inter -cross (MAGIC) population was subjected to high -throughput chlorophyll fluorescence phenotyping under fluctuating light to determine time -dependent quantitative trail loci (QTL) for photosynthetic regulation. QTL were identified tha t showed a time-dependent emergence and disappearance relative to fluctuations in light intensity. Utilizing this high -throughput phenotyping approach combined with the identified QTL, alterations in the onset and dissipation of NPQ could be manipulated to improve plant productivity in field conditions. 132 5.2 Introduction Increases in crop productivity during the 20 th century, the so -called Green Revolution, were driven primarily by increases in the partitioning of biomass into grain (harvest index) (190). While improvements in biomass accumulation depend upon the initial utilization of light by photosynthesis, increases in the efficiency of photosynthesis have not necessarily been improved with breeding for improved harvest index (191). Meeting the inc reased global demand for food by 2050 (192) will require substantial gains in yield per hectare; with crop breeding producing plants that are near the theoretical upper limit in harvest index (193, 194), improvements in photosynthetic efficiency, which is far from the theoretical biological limit (103, 195), are promising avenues for the improvement of crops (196). Under natural field conditions, plants experience a variety of biotic and abiotic factors that influence their overall productivity. Photosynt hetic light reactions, although dependent on light availability for the production of cellular energy, are sensitive to the abrupt fluctuations in light intensity experienced in nature from cloud cover and other physical shading that lead to rapid changes in the incident light intensity upon a leaf (60, 66, 75, 197). These light intensity fluctuations have been shown impact the yield of plants relative to laboratory conditions under which the basic processes of photosynthesis are usually studied (66), neces sitating an improvement of our understanding of the operation and regulation of the light reactions under natural, environmental light conditions. Oxygenic photosynthesis in cyanobacteria, algae, and plants utilizes multiple membrane Ðlocalized protein co mplexes in series to mediate the overall transfer of electrons from water to ferredoxin and NADPH (127). The electron transfer processes of photosynthesis are stabilized and coupled to the transfer of protons into the thylakoid lumen, generating a light -induced proton 133 gradient ( ! pH) across the thylakoid membrane which is utilized by the thylakoid ATP synthase to generate ATP (76). As the thylakoid lumen pH decreases, protonation of lumen -exposed residues of specific proteins leads to feedback regulation of the light reactions (61). Quinol oxidation by cytochrome b6f decreases 10 -fold per pH unit decrease, which slows electron transfer from photosystem II (PSII) (11, 12). Thermal dissipation of absorbed light as heat, termed nonphotochemical quenching (NPQ), is initiated via protonation of violaxanthin de -epoxidase (VDE) (62) and protonation of the thylakoid membrane protein PsbS (7). The activation of these NPQ processes decreases the excitation pressure on PSII, preventing the over -reduction of PSII electron acceptors (6). While the ! pH component of the thylakoid proton motive force ( pmf ) has well -established roles in regulating photosynthesis, the electric field ( !!) component of the pmf has emerged as a regulator of electron transfer reactions in vivo as well (36). As the thylakoid !! increases, the vectorial transfer of electrons within protein complexes becomes increasingly thermodynamically destabilized (38), leadi ng to an increase in reverse electron transfer reactions (i.e. recombination), which can result in the production of reactive oxygen species (ROS) and damage to the photosystems resulting in photoinhibition (56). While NPQ results in photoprotection of PSI I by decreasing the excitation pressure, the kinetics of ! pH generation are limited by the high buffering capacity of the thylakoid lumen (22), while !! generation kinetics are much more rapid due to the low electrical capacitance of the membrane (20). The se kinetic imbalances result in short periods where !! dominates the pmf composition prior to ! pH buildup, which in turn leads to a kinetic lag prior to NPQ activation where PSII is subject to high excitation pressure as well as increased probability of el ectron recombination and ROS production (167). 134 While the kinetics of NPQ induction may limit plant productivity by increasing the probability of !!-induced photoinhibition, the NPQ dissipation kinetics have also been suggested to limit plant productivity (125, 198, 199). Upon a decrease in light intensity, slow relaxation of NPQ can limit productivity by sustaining exciton quenching in the antenna and limiting PSII excitation under the now low -light condition, leading to less net photosynthesis during the time required to dissipate the quenching state. Genetic engineering of NPQ has proven one potential avenue to circumvent these losses and improve biomass accumulation (126). The dramatic impacts that the kinetic mismatches between light intensity changes and NPQ induction/dissipation impose on plant productivity may have lead, over evolutionary time, to divergence in the kinetics of NPQ induction and deactivation. Variation in the overall capacity for NPQ exists between species (200, 201), between shade v s. high -light grown plants (201, 202), and within a species (203-205). However, while natural variation in photosynthetic parameters has been observed within different genetic accessions of the same species, a correlation between maximum rates of photosyn thesis (measured under saturating light) does not necessarily correlate positively with biomass accumulation (206-208). Although the accumulation of biomass by photosynthetic organisms begins with the transformation of light into chemical energy, the multi faceted regulatory processes determining the partitioning of accumulated biomass could be one explanation for inconsistent or lack of correlation between photosynthesis and biomass accumulation (209). However it is also possible that the technical challeng es of measuring photosynthesis under non -static conditions have limited the potential identification of genetic factors influencing photosynthesis under more natural conditions. To overcome this hurdle, we have utilized chlorophyll fluorescence imaging und er dynamic environmental conditions (75) in combination with an Arabidopsis thaliana multi -parent 135 advanced generation inter -cross (MAGIC) population (210) to map natural variation in photosynthesis during light fluctuations. This approach allows photosynth etic measurements to be obtained simultaneously for dozens of plants under non -static conditions that can be precisely manipulated to rapidly phenotype plants under environmental conditions. 5.3 Materials and Methods 5.3.1 Plant materials and growth cond itions The MAGIC lines (210) as well as the 19 parental accessions were obtained from the Arabidopsis Biological Resource Center (http://abrc.osu.edu). Freshly propagated seed was generated for each genotype prior to phenotyping experiments. Plants were gr own on soil at 100 µmol photons m -2s-1 for a 16 hr/8 hr light/dark photoperiod at 22¡C. 5.3.2 Chlorophyll fluorescence imaging Whole -plant chlorophyll fluorescence imaging was performed on plants 21 days after germination in custom outfitted growth chambe rs (75). To minimize any potential phenotypic variation due to any differences in circadian rhythm, all plants were imaged during the first hour of illumination during the photoperiod. During the imaging procedure, the light intensity was maintained for 10 min at each of five intensities: 100, 600, 200, 1000, and 100 µmol photons m -2s-1 (Fig. 5.1). For high -density measurements of steady -state fluorescence ( "f), Fo was measured after 8 hours of dark adaptation corresponding to the ÒnightÓ portion of the pho toperiod. At the start of the light portion of the photoperiod, measurements of "f were obtained every 30 s over the course of illumination. The fluorescence yield in the light ( "fs) at each point was normalized to the minimum fluorescence yield in the dar k Fo. Variations in quantitative trait loci ( QTL) identification did not occur with or without normalization to F o. 136 For measurements of photosynthetic parameters derived from pulse amplitude modulation, F V/FM was measured 30 s prior to the onset of actinic illumination after 8 hours of dark adaptation corresponding to the ÒnightÓ portion of the photoperiod. Upon actinic illumination, chlorophyll fluorescence images were acquired every 2 min during illumination using saturation pulse methods (75, 211). At least three replicates for each genotype were measured for each of the photosynthetic parameters was used for QTL analysis. Due to lack of germination or survival until the measuring stage, some MAGIC genotype s did not have at least three biological replicates, and the genotypes were removed from subsequent QTL analysis. Fluorescence images were analyzed using open source image analysis software (ImageJ, NIH) modified in house to calculate photosynthetic parame ters over whole plant averages. 5.3.3 QTL analysis QTL analysis was performed using the HAPPY R software package as in Kover et al. (210). Briefly, this utilizes experimentally tested markers and a genetic map to construct a mosaic genome for each MAGIC g enotype. Using a probabilistic hidden Markov model, the founding genotype for each of 1,260 single nucleotide variations (SNPs) is tested and determined for each genotype. The likelihood of no QTL being present at each phenotype was tested by fitting a fix ed-effect linear model corresponding to a standard genome scan with up to 18 degrees of freedom and performing an analysis of variance (ANOVA). The statistical significance for each locus was summarized by the negative logarithm of the ANOVA p-value ( -log 10 p). The genome -wide significance thresholds were independently determined for each measurement time point by repeating the genome scan with 1000 permutations. To minimize the identification of 137 false positive QTL, a QTL was only defined if the Ðlog 10p val ue of the locus was above the permutation derived 95% genome -wide significance threshold. 5.3.4 Protein extraction and western blotting Analysis of proteins with known involvement in NPQ was performed using total leaf protein from each of the 19 Arabidops is thaliana MAGIC founder accessions as in Livingston et al. (118). To minimize variation in protein levels due to developmental differences between leaves (212), and due to the variation in number of rosette leaves between the different accessions, protei ns were extracted from three pooled biological replicates of the youngest fully expanded leaf of 21 day -old plants. Determination of specific proteins was performed by separating 20 µg of total leaf proteins via SDS -PAGE under reducing conditions followed by transfer to polyvinylidene difluoride (PVDF) membrane. NPQ proteins were analyzed using antibodies specific to each protein, anti -violaxanthin de -epoxidase ( $-VDE, PHY1225S, PhytoAB, Redwood City, CA, USA), anti -zeaxanthin epoxidase ( $-ZEP, PHY0499, Phy toAB, Redwood City, CA, USA), and anti -PsbS ( $-PsbS, AS09 533, Agrisera, Vannas, Sweden) and detected using an anti -rabbit secondary antibody conjugated to alkaline phosphatase. Samples were developed via addition of BCIP/NBT substrates (Sigma -Aldrich, St. Louis, MO, USA). 5.4 Results 5.4.1 Variation in NPQ kinetics between Arabidopsis accessions To discover underlying factors regulating photosynthesis under conditions mimicking natural field stresses, NPQ was assayed over time during the course of fluctuating light in the 19 Arabidopsis thaliana founding accessions used to create the Arabidopsis MAGIC population (Fig. 5.1) (210, 213). Rather than compare the extent of NPQ in low light versus high light, the dynamics of NPQ changes during fluctuating light was used, as the rapid induction of NPQ 138 following an increase in irradiance is proposed to de crease the probability of PSII photodamage (167) while the rate of NPQ relaxation when the irradiance decreases will alter the capability of PSII to utilize the non -inhibitory light (126). During light fluctuations (Fig. 5.1A), the 19 MAGIC founders displa y a range of NPQ phenotypes that represent all three potential scenarios for NPQ regulation: fast vs. slow induction, fast vs. slow dissipation, and altered steady -state NPQ extent (Fig. 5.1B, C). Figure 5.1: The 19 founding parental accessions of the Arabidopsis MAGIC population display a range of NPQ induction and dissipation kinetics during fluctuating light. Chlorophyll fluorescence imaging of NPQ in the 19 founding Arabidopsis thaliana accessions of the MAGIC population was carried out during fluct uations in light intensity (A). Light intensities were: 100, 600, 200, 1000, and 100 µmol photons m -2s-1. The light was maintained at each intensity for 10 min. Following dark -adaptation corresponding to the normal ÒnightÓ portion of the photoperiod, the m aximum quantum yield of PSII photochemistry was measured to obtain FM. The imaging period corresponded to the first hour of light during the photoperiod, with the first 5 measurements (100 µmol photons m -2s-1) occurring at growth light intensity. NPQ was measured every 2 min in the light (B) averaged over whole plant rosettes (C). For visualization, A B Col-0 Ct-1 Rsch-4 C 120 240 360 480 600 720 840 960 1080 1200 1320 1440 1560 1680 1800 1920 2040 2160 2280 2400 2520 2640 2760 2880 3000 0 3.2 Time (s) 139 Figure 5.1 ( continued ): standard deviation is not shown in B (mean, n = 6-8). False -color NPQ images of representative plants of 3 accessions (Col -0, Ct-1, Rsch -4) over the course of the imaging procedure (C) show time -dependent changes in NPQ as well as total NPQ differences. To investigate potential causes of NPQ variation among the founding genotypes, differences in the key regulators of the pH -dependent component of NPQ (q E) were assessed (Fig. 5.2). Alterations in PsbS or VDE within the population could explain differences in the ability of plants to induce NPQ (8), while alterations to zeaxanthin epoxidase (ZEP) to relieve xanthophyll dependent qE quenc hing component could be a factor in dissipating NPQ at low light (166). Among the 19 founding accessions, the protein content of PsbS is approximately the same among each plant (Fig. 5.2C), suggesting that differences in NPQ extents during the fluctuating light experiment is not due to alteration of PsbS protein levels. However, enzymes responsible for the conversion of xanthophyll pigments do show differences in the level of protein con tent relative to total leaf protein (Fig. 5.2A, B). Although measured at different developmental stages (21 day -old plants in current work versus ~12 day -old plants in Gan et al.), Gan et al. also found that the transcripts for VDE ( AT1G08550 ) and ZEP ( AT5G67030 ) were differentially expressed within the founding accessions, suggesting that regulation of the VDE/ZEP ratio in vivo represents at least one strategy that natural populations have evolved to regulate NPQ (47). Contrary to RIL analysis where phenot ypic differences require binary parental differences, most of the founding accessions show no differences in the protein levels of any of the three components, but still have altered NPQ dynamics relative to the other accessions, indicating that multiple f actors influence the extent and rates of NPQ and will be present in the MAGIC population. 140 Figure 5.2: Differential content of proteins mediating the rapidly reversible pH -dependent qE component of NPQ Arabidopsis accessions. Total leaf protein was extr acted from 21 day -old plants for each Arabidopsis founding accession. Protein levels for ZEP (A), VDE (B), and PsbS (C) were determined from 20 µg of total leaf protein using antibodies specific to each protein. Coomassie staining of 20 µg of the same protein samples (D) was performed to ensure equal loading across samples. 5.4.2 QTL mapping of NPQ kinetics To elucidate factors influencing photosynthetic responses to fluctuating light, 379 MAGIC genotypes were measured under the same conditions as the fo unding accessions (Fig. 5.3). Similar to the founders, different dynamics of NPQ induction, dissipation, and total extent were present across the phenotypes of the MAGIC population. QTL analysis of NPQ identified 71 QTLs above the genome -wide 95% significa nce threshold at 19 of the measurement times (Fig. 5.4, Table 5.1). Over the entire experimental period, this represents 21 unique QTL identifications, as the same QTL could be identified multiple times at different measurements. During the measuring perio d, the photochemical yield of PSII ( #II) was determined at the same time as NPQ, however no QTLs were observed for #II. Transformation of the #II data to estimate linear electron flow ( #II * PAR * PSII cross -section * A), estimating a PSII cross -section of 0 .5 and leaf absorptivity of 0.8 for all plants, also yielded no QTL. Future analyses measuring the Bur-0 37 50 !-ZEP !-PsbS Can-0 Col-0 Ct-1 Edi-0 Hi-0 Kn-0 Ler-0 Mt-0 No-0 Oy-0 Po-0 Rsch-4 Sf-2 Tsu-0 Wil-2 Ws-0 Wu-0 Zu-0 A B C D !-VDE 141 specific light absorption parameters for each MAGIC genotype could better define QTL related specifically to PSII photochemistry. Figure 5.3: Kinetics of NPQ responses in the Arabidopsis MAGIC population under fluctuating light. Whole plant chlorophyll fluorescence imaging was used to phenotype the NPQ responses of the MAGIC population over a fluctuating light time course. The mean NPQ at each time point of n = 3-4 biological replicates of each MAGIC genotype is clustered by the total phenotype over the course of the experiment. The position of each founding accession is indicated by the colored bar next to the dendrogram. Accession colors are the same as in Fig. 5.1. 0 500 1000 PAR 142 Figure 5.4: Time -dependent identification of NPQ QTL during fluctuating light in Arabidopsis MAGIC lines. QTL analysis of NPQ phenotypes of the MAGIC population were determined every 2 min during a fluctuating light treatment, resulting in 5 NPQ measurements per light intensity. The peak positions of QTLs with Ðlog 10p-values > 95% genome -wide significance are shown relative to the position in the Arabidopsis thaliana Col-0 reference genome at the time at which they are significant until the next measured time point. Vertical dashed lines indicate the changes in light intensity. 5.4.3 QTL mapping of rapid responses to light fluctuations Utilizing pulse amplitude modulated (PAM) fluorometry, the extent of different photosynthetic proces ses can be determined (211). However, the high intensity of light used during the saturating pulses to fully reduce PSII limits how rapidly the pulses can be applied; as the interval between saturation pulses decreases, damage to photosynthetic proteins be gins to occur, decreasing the ability to measure how rapidly processes change following a disturbance. Another limitation of using chlorophyll fluorescence as a probe of photosynthesis is that information about the function of PSII is obtained while other processes of the light reactions, 143 pmf extent and composition, b6f turnover, PSI redox state, and ATP synthase activity, are not directly observed. To increase the capacity to measure rapid changes in photosynthesis, the yield of chlorophyll fluorescence ( "f) was measured every 30 s during the fluctuating light regime separately from experiments measuring NPQ. The "f is determined by the rate constant of fluorescence relative to the rate constants for all of the processes that compete for excitation energy: !!!!!!!!!!!!"!!!!"!!!!"!!!!"!!!!!!"#!!!!!!!! (Eq. 1) where kf is the rate of fluorescence, kqE the rate of energy -dependent nonphotochemical quenching q E, kqE the rate of photoinhibitor y nonphotochemical quenching q I, kqT the rate of state transition nonphotochemical quenching, kisc the rate of non -radiative decay due to intersystem crossing, kd the rate of non -radiative decay, kpc the rate of photochemistry, and q L the fraction of PSII open reaction centers (PSII with oxidized Q A). In higher plants, quenching due to antenna state transitions appears to be minimal (155). kqE is a composite rate of the various processes that mediate the total q E response, the rates of pH -regulated PsbS que nching, pH -mediated activation of VDE and conversion of violaxanthin into zeaxanthin, and the activity of zeaxanthin epoxidase to convert zeaxanthin to violaxanthin (146). The redox state of Q A (qL) is also regulated by pH -dependent downregulation of cytoc hrome b6f turnover, increasing the fraction of Q A- as the thylakoid lumen pH decreases. The photosynthetic pmf is composed of both a ! pH and !! (2, 39, 135, 137), with the partitioning regulated by the activity of thylakoid ion channels (34, 35). While th e ! pH has a well-studied role in regulating photosynthesis to limit photodamage, a large !!, which occurs following light fluctuations to a higher light intensity, has been shown to promote photodamage 144 through reactive oxygen species production (36). There fore the kinetics of !! dissipation and ! pH generation to mediate photosynthetic regulation following light transitions could be a limiting factor for plant productivity (167). Therefore, although the photosynthetic pmf cannot be imaged directly, changes i n "f following light fluctuations will respond proportionally to changes in ! pH generation or relaxation due to the direct regulation of quenching processes by the lumen pH. To follow the response of "f to light fluctuations, "f was imaged every 30 s d uring the fluctuating light treatment (Fig. 5.5) in 422 MAGIC genotypes. To differentiate "f variation due to time Ðdependent kinetic changes due to light fluctuations from intrinsic variations in fluorescence yield which could have shifted amplitudes but identical kinetics, "f was normalized to the first measurement after each light intensity shift to determine how rapidly each genotype responded to changes in light intensity. Changes in "f did not follow a simple ex ponential decay over time, limiting the ability to fit the "f response of each genotype and utilize the corresponding decay rate as a phenotype for QTL analysis. Instead, the amplitude of "f change from the normalized t0 of each light change was determine d (Fig. 5.5A), and this phenotype used to determine time -dependent QTL (Fig. 5.5C). As the composition of the thylakoid ! pH changes following a light fluctuation, the dual regulation of q E and b6f turnover mediates the concentration of PSII with Q A-, which is reflected in the amplitude of "f - "f t0, with how rapidly "f changes from the light fluctuation proportionally reflecting how rapidly ! pH changes. 145 Figure 5.5: The kinetics and amplitude of chlorophyll fluorescence quenching varies among the Arabido psis MAGIC founders and MAGIC population during light fluctuations. Using the same light intensity changes as the NPQ measurements (Fig. 5.1) chlorophyll fluorescence ( "f) was measured every 30 s in the Arabidopsis founding accessions (A). To quantify the changes in "f over time, each "f measurement was normalized to the first measurement within each light intensity. Variation in the phenotypes within the MAGIC population (B) for the change in "f show broad distributions from the mean of the population as well as shifts in the mean of the population upon transitions from high light to low light. The mean of each founding accession is indicated by colored symbols above each histogram. The mean "f - "f t0 at each time point of n = 3-4 biological replicates of each MAGIC genotype is clustered by the total phenotype over the course of the experiment (C). The position of each founding accession is indicated by the colored bar next to the dendrogram. For visualization, the first measurement at each light intensity, used for normalizing, are not shown (all values = 1). Colors of each founding accession are maintained throughout each of A, B, and C. Analysis of the variation in the change in "f over time identified 129 QTL above the genome -wide 95% significance thr eshold (Fig. 5.6, Table 5.2). Over the course of the fluctuating light regime, multiple instances occur of a QTL centered at the same locus, narrowing the total 0!100 100!600 1000!100 µmol m -2s-1 600!200 200!1000 A B C Time from light change 30s 10 min 146 number of QTL to 57 unique QTL peaks. Surprisingly, most of the identified unique QTL (50/57) were only identified upon the r ecovery from high light to low light. Following transitions from lower light intensity to high light, only seven unique QTL peaks were identified. While increases in light intensity were presumed to lead to the greatest changes in the "f relative to the initial values, a possible explanation for the large differences between the conditions leading to QTL identification is the variation in mean phenotypes within the MAGIC population under these conditions. The distribution of phenotypes over time is less di vergent from the mean following low light to high light fluctuations than the transition from high light to low light (Fig. 5.5B). During the initial induction of photosynthesis from overnight dark adaptation, an early responding QTL centered at 5.14 Mb on chromosome 1 was identified at 90 -180 s following the beginning of the photoperiod, with a late responding photosynthetic induction QTL identified at 9-9.5 min after the beginning of the photoperiod, potentially identifying QTL representing constitutive p hotosynthetic differences between the genotypes. The transition to 1,000 µmol photons m -2s-1, above the normal light saturation for Arabidopsis, identified five closely localized QTL on chromosome 1, cumulatively explaining ~10 -15% of the total phenotypic variance at each time point. 147 Figure 5.6: Time -dependent changes in the amplitude of "f quenching during light fluctuations. To determine how rapidly photosynthesis responds to changes in light intensity, QTL analysis was performed on the amplitude of "f every 30 s relative to the first "f measurement at each light intensity. The peak positions of QTLs with Ðlog 10 p-values > 95% genome -wide significance are shown relative to the position in the Arabidopsis thaliana Col-0 reference genome at the time at which they are significant until the next measured time point. Vertical dashed lines indicate the times at which the light intensity changes. 5.5 Discussion The utility of Arabidopsis thaliana as a genetic model allows many phenotypes to be analyzed and understood at the genetic level (214). The major contributors regulating the light reactions of photosynthesis in the green lineage have been well characterized at the molecular level in vitro and in vivo , with mutants in Arabidopsis thaliana and Chlamydomonas reinhardtii identifying many of the specific proteins involved in pH -mediated regulation of photosynthesis (reviewed in 5). However, while analysis of knockout mutants for specific genes cl early identifies the role of that gene in physiological processes, the in vivo regulation in terms on/off 148 rates, total extent, and changes due to acclimation are lost when using only knockouts to understand the regulation (215). Under most laboratory condi tions, where plants are undergoing minimal if any stress, the regulatory significance underpinning physiological processes can be lost. While laboratory conditions have helped dissect many of the molecular mechanisms of NPQ regulation of photosynthesis (166, 176, 216), understanding the regulation of photosynthesis in environmental conditions requires further progress (60, 75). The regulation of excitation pressure reaching PSII complexes within leaves is mediated by feedback regulation from the light -gene rated ! pH, modulating the redox state PSII electron acceptors by decreasing excitation energy transfer via q E activation and decreasing electron transfer via regulation of b6f quinol oxidation (5). These processes help minimize the amount of PSII photodama ge, which irreversibly inhibits the damaged PSII complex and requires degradation and de novo synthesis of a new D1 subunit to maintain photosynthetic integrity (128). While protease activities and de novo protein synthesis have an energetic (ATP) cost, of larger concern to plant productivity is the decreased photosynthetic capacity that occurs while the PSII complex is inactive (65). Therefore, while downregulation of photosynthesis via NPQ and electron transfer regulation limit the output of photosynthesi s, the evolution of these photoprotective processes and distribution among photosynthetic organisms suggests that the benefits of dampening photosynthesis outweigh the negative growth productivity due to enhanced photodamage (66). The protection of photos ynthesis under high light intensities, however, limits photosynthetic capabilities as soon as the light intensity decreases (195, 199). Conversely, due to kinetic limitations that limit the buildup of ! pH versus !! (2), slow induction of q E is suggested to increase PSII photodamage (36, 167). Taken together, improvements in overall photosynthetic 149 yields could be accomplished by increasing the rate at which q E induction occurs while simultaneously increasing q E dissipation when it is no longer necessary. To identify factors that would allow both of these conditions to occur, variation in Arabidopsis thaliana natural populations was utilized to identify QTL involved in rapid photosynthetic responses to light fluctuations. Although many of the core genes of th e photosynthetic reaction centers are encoded by the chloroplast genome, natural diversity of nuclear genomes has been utilized to improve the understanding of photosynthetic regulation and factors that are involved in modulating the basic bioenergetics pr ocesses of the light reactions (203, 204, 217). Whole plant chlorophyll fluorescence phenotypes for both NPQ as well as changes in "f quenching identified time -dependent fluctuating light QTL in the Arabidopsis MAGIC population (Figs. 5.4, 5.6). While vari ations in NPQ leading to the identification of QTL were present at most time points (Fig. 5.4), changes in "f relative to the light intensity fluctuations primarily appear after high light to low light transitions (Fig. 5.6). The time -dependence of these QTL relative to when the light fluctuates is most apparent for how rapidly "f quenching changes. After a light fluctuation from 600 to 200 µmol photons m -2s-1, early QTL on chromosomes 1, 2, 4, and 5 appear within 30 s of t he light intensity change and are not longer present 90 s after the light decrease, while late QTL on chromosomes 3 and 5 emerge after ~3 minutes (Fig. 5.6). Using a multi -parent population, as was done in the present work, the increased nucleotide poly morphism and crossover frequency obtained using genome -wide association panels is approached, while maintaining limited genome structure complexity provided by recombinant inbred lines (RIL) (210, 213). As such, within the MAGIC founding accessions, variat ion in NPQ processes is likely mediated in part by cis-regulation of the protein levels of 150 VDE and ZEP (Fig. 5.2) in some founding accessions as well as the progeny. Altering the ratio of VDE/ZEP has been suggested as a strategy to alter the apparent pH -dependency of q E activation (47), allowing rapid q E activation with decreased ZEP levels such as is seen in Wu -0 (Figs. 5.1, 5.2). Further analysis of xanthophyll pigment compositions and pigment changes over time will help clarify how much the altered prote in contents mediate NPQ phenotypic variation. Similar to other analyses of natural populations, variation in PsbS protein content does not explain the variation in NPQ within the founding accessions (205, 215). Advantageously, relative to a RIL population, using a multi -parental mapping population the phenotypes are not limited to binary phenotypes of the two parental lines used to generate a RIL population. As such, although VDE/ZEP levels may be one route used to modulate NPQ dynamics in Arabidopsis, foun ding accessions with no apparent differences in the level of NPQ component proteins still display a range of NPQ phenotypes (e.g. Ct -1 vs. Rsch -4 in Fig. 5.1), suggesting that other factors regulating NPQ could be identifiable within the QTL. The identifi cation of photosynthetic QTL in a time -dependent manner during fluctuating light conditions helps shift the strategies for identifying mechanisms utilized to maintain photosynthesis under natural conditions (197). Although genomic variation within the foun ding accessions of the MAGIC population is similar to the global Arabidopsis nucleotide diversity, polymorphisms that generate amino acid changes in protein coding genes are underrepresented (213). Within the MAGIC founders, 80% of the predicted proteins h ave major isoform frequencies of at least 15/19, meaning that each polymorphism causing a functional protein change is likely to be represented in 4 or fewer of the 19 lines, effectively recapitulating RIL population structures of bi -allelic changes at mos t loci conferring function protein changes. Further refinement of the QTL mapping regions combined with candidate allele testing will help 151 provide evidence for alleles within the population contributing to rapid photosynthesis responses to light fluctuatio ns. These can subsequently be tested for their impact on yield under field conditions (126) as well as used in marker -assisted breeding in crop species to boost the initial steps of plant anabolism in order to increase yield (218). Incremental improvements in the initial steps of photosynthesis under dynamic light conditions facilitated by the rapid, high -throughput phenotyping utilized in this study, combined with genetic identification and implementation of beneficial alleles, will help drive increases in plant productivity. 5.6 Acknowledgements We would like to thank Linda Savage for initial discussions on planting strategy for screening the MAGIC population. Work performed by G.A.D. and D.M.K. was supported by the US Department of Energy (DOE), Office of Science, Basic Energy Sciences (BES) under award number DE -FG02 -91ER20021. 152 Table 5.1: List of QTLs identified for time -dependent NPQ responses to fluctuating light. Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 120 100 120 2902408 4 2502452 3006991 3.76 120 100 120 6647145 2 6154830 6647145 3.60 120 100 120 23247571 5 23246477 23248880 3.60 360 100 360 15047966 5 14694836 15439873 4.38 480 100 480 15047966 5 13832746 15750717 4.63 600 100 600 13201153 1 11850168 13832436 4.49 600 100 600 15047966 5 13832746 15439873 4.28 720 600 120 13090494 1 11399904 17679130 6.88 720 600 120 14935674 5 12512173 16337385 6.40 720 600 120 21864028 5 19397857 23246248 5.64 720 600 120 18166800 5 17341215 18765193 4.43 720 600 120 11067093 5 10525858 12183470 4.00 720 600 120 6025716 2 5666979 6647145 3.94 840 600 240 22023651 5 17538410 23247571 7.26 840 600 240 13090494 1 11399904 17876791 6.92 840 600 240 15298812 5 12426277 16429063 6.61 840 600 240 23798428 5 23705688 23810485 4.32 840 600 240 6025716 2 5666979 6647145 4.18 960 600 360 22023651 5 19106920 23247571 7.01 960 600 360 15048131 5 12576814 16337385 6.42 960 600 360 13090494 1 11412672 17679130 5.83 960 600 360 23798428 5 23705688 23810485 4.27 1080 600 480 22023651 5 18266506 23247571 7.02 1080 600 480 15298812 5 12576814 16429063 6.55 1080 600 480 13090494 1 11412672 16134927 5.74 1080 600 480 16872100 1 16251782 17679130 4.48 1080 600 480 23798428 5 23559674 23810485 4.49 1080 600 480 18166800 5 17461027 18241539 3.99 1200 600 600 15298812 5 12681159 16429063 6.36 1200 600 600 22023651 5 18278591 23247571 6.78 1200 600 600 13090494 1 11466186 15525445 5.12 1200 600 600 23798428 5 23559674 23810485 4.25 1920 1000 120 13201153 1 11037687 17876791 7.87 1920 1000 120 15048131 5 12512173 16337385 5.34 1920 1000 120 22023651 5 21021855 22610276 4.96 1920 1000 120 8978323 5 8360127 9481360 4.09 153 Table 5.1 (continued ): Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 1920 1000 120 23098340 5 22697296 23246477 3.67 2040 1000 240 13201153 1 11070732 17876791 8.34 2040 1000 240 15048131 5 12512173 16186076 5.36 2040 1000 240 22023651 5 21021921 22610276 4.92 2040 1000 240 8978323 5 0 9481360 4.18 2040 1000 240 11067093 5 10525702 12183470 3.90 2040 1000 240 23098340 5 22697296 23246477 3.79 2160 1000 360 13201153 1 11159782 17876791 8.16 2160 1000 360 15048131 5 12681159 16186076 4.90 2160 1000 360 22023651 5 21243342 22610276 4.56 2160 1000 360 8978323 5 0 9481360 3.91 2160 1000 360 11067093 5 10525858 11613391 3.61 2160 1000 360 23246248 5 22697296 23246477 3.58 2280 1000 480 13201153 1 11159782 17876791 7.97 2280 1000 480 15048131 5 12681159 15750933 4.79 2280 1000 480 22023651 5 21701553 22397775 4.48 2280 1000 480 8978323 5 0 9481360 3.80 2280 1000 480 21398449 5 21243342 21398449 3.61 2280 1000 480 23246248 5 22697296 23246477 3.59 2280 1000 480 11067093 5 10525858 11613391 3.54 2400 1000 600 13201153 1 11399904 17876791 7.53 2400 1000 600 15048131 5 12681159 15750933 4.76 2400 1000 600 22023651 5 21701553 22397775 4.39 2400 1000 600 8978323 5 8810481 9481360 3.72 2520 100 120 13540607 1 12892642 13832436 3.81 2640 100 240 13832180 1 12208016 13832436 4.28 2640 100 240 15298812 5 14935674 15439873 3.77 2760 100 360 13832180 1 12208016 14160452 4.52 2760 100 360 15298812 5 14694836 15439873 3.98 2880 100 480 13201153 1 11850168 16128552 4.82 2880 100 480 15298812 5 13832746 15439873 4.36 3000 100 600 13201153 1 12208016 14160452 4.46 3000 100 600 15298812 5 13832746 15439873 4.06 154 Table 5.2: List of QTLs identified for time -dependent changes in "f amplitude. !Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 90 100 90 5148226 1 5095611 5206772 3.66 120 100 120 5148226 1 4992111 5206986 3.81 150 100 150 5148226 1 4947328 5290747 3.99 180 100 180 5148226 1 4992111 5206986 3.73 570 100 570 13201153 1 12208016 13201153 3.64 600 100 600 13201153 1 12208016 13201153 3.76 930 600 330 6416383 5 6415440 6708110 3.59 960 600 360 6416383 5 6222069 6899955 3.91 960 600 360 5996219 5 5829162 5996219 3.62 1290 200 90 2902408 4 2695395 4197470 4.66 1290 200 90 10423538 1 8974265 10841461 4.38 1290 200 90 1585202 4 1243348 2510399 4.39 1290 200 90 23798306 5 23300895 24816019 3.84 1290 200 90 11412672 1 11159782 12093566 3.67 1290 200 90 21701553 5 21398449 21701553 3.68 1290 200 90 11268000 4 10659145 11470264 3.62 1320 200 120 13979309 2 13319028 15244621 4.40 1320 200 120 13090494 1 11655539 13217139 3.67 1320 200 120 1553591 4 1243348 1585202 3.65 1320 200 120 10423538 1 8974265 10547614 3.50 1320 200 120 11412672 1 11215275 11553534 3.50 1380 200 180 4073395 3 3679535 4073911 3.93 1410 200 210 24295906 5 23903662 24816019 3.50 1440 200 240 8541646 5 7315346 8978323 3.87 1440 200 240 4073911 3 3991976 4073911 3.78 1500 200 300 4073911 3 3679286 4073911 4.06 1500 200 300 3344784 3 2903318 3610515 3.93 1650 200 450 3344784 3 2903318 3580015 4.02 1710 200 510 11466186 1 11159852 11850168 3.49 1800 200 600 11215275 1 11159782 11553534 3.66 1800 200 600 8810481 5 8001188 9628385 3.63 1860 1000 60 297887 3 276915 819034 4.08 1860 1000 60 965025 3 819454 965025 4.08 1890 1000 90 20135840 1 19502363 20310446 3.99 1920 1000 120 20135840 1 19502363 20510777 4.54 155 Table 5.2 (continued ): Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 1920 1000 120 17679130 1 17148697 19434966 3.91 1980 1000 180 20135840 1 19502363 20310446 4.00 1980 1000 180 16871886 1 16134927 16872100 3.74 2040 1000 240 20135840 1 19778790 20171160 3.60 2160 1000 360 20135840 1 19778790 20310446 3.65 2280 1000 480 20135840 1 19502363 20310446 3.94 2280 1000 480 16871886 1 16134927 17179544 3.82 2280 1000 480 18943964 1 17474215 19397107 3.67 2310 1000 510 20135840 1 19502363 20310446 4.06 2310 1000 510 16871886 1 16134927 16872100 3.80 2310 1000 510 18629407 1 17474215 19434966 3.74 2340 1000 540 20135840 1 19502363 20310446 3.97 2370 1000 570 20135840 1 19502363 20310446 3.81 2400 1000 600 16871886 1 16134927 16872100 3.66 2400 1000 600 20135840 1 19778790 20310446 3.64 2460 100 60 11268000 4 9952391 11786400 4.46 2460 100 60 13832746 5 12681159 15439873 4.23 2460 100 60 10525702 5 9628385 10910094 3.86 2490 100 90 11268000 4 9199369 13576430 6.97 2490 100 90 7722418 2 7544501 7722418 4.33 2490 100 90 13832746 5 13597330 17130805 4.09 2520 100 120 11268000 4 9199369 13670757 7.88 2520 100 120 16351105 5 14973938 17907459 4.14 2520 100 120 7722418 2 7643146 7722418 3.94 2520 100 120 9579136 2 9249015 10127320 3.70 2550 100 150 11268000 4 9199369 13670757 7.07 2550 100 150 16947516 5 15744761 18048533 3.93 2550 100 150 2902408 4 2803493 2907158 3.80 2580 100 180 11577769 4 9630919 13078462 6.48 2580 100 180 10127320 2 9249015 11142985 3.92 2580 100 180 2902408 4 2803493 3001617 3.85 2610 100 210 11268000 4 9630919 12900417 6.25 2610 100 210 2902408 4 2803493 3001617 3.75 2610 100 210 10423538 1 9973305 10720291 3.67 156 Table 5.2 (continued ): Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 2610 100 210 22606285 3 21773266 22627912 3.65 2610 100 210 23248880 5 23247571 23248880 3.45 2640 100 240 11577769 4 9630919 12900417 5.26 2640 100 240 2902408 4 2803493 3129542 4.13 2670 100 270 11577769 4 9952391 11984761 4.70 2670 100 270 2902408 4 2695395 3924075 4.59 2670 100 270 23705688 5 23253768 24357567 3.94 2670 100 270 23247571 5 23246477 23248880 3.64 2670 100 270 1585202 4 1243079 1585202 3.63 2700 100 300 2902408 4 2695395 3924075 4.75 2700 100 300 11580131 4 9952391 11878383 4.36 2700 100 300 23798306 5 23253768 24357567 3.90 2700 100 300 1585202 4 1241726 1585202 3.70 2700 100 300 1132454 4 1125812 1132454 3.63 2730 100 330 2907158 4 2803493 3924075 4.07 2730 100 330 11580131 4 9952391 11878383 4.01 2730 100 330 23300895 5 23246477 24295906 3.96 2730 100 330 10423538 1 9516375 10720291 3.57 2760 100 360 2902408 4 2695395 4253906 4.63 2760 100 360 10777260 4 9952391 11878383 4.41 2760 100 360 23798428 5 23253768 24357567 3.87 2760 100 360 23247571 5 23246477 23247571 3.76 2760 100 360 1585202 4 1241726 1585202 3.61 2760 100 360 1585202 4 1241726 1585202 3.61 2760 100 360 3991976 3 2967872 4073911 3.56 2760 100 360 1132454 4 1125812 1237935 3.56 2790 100 390 10777260 4 9952391 11822736 4.44 2790 100 390 2902408 4 2803493 3535626 3.84 2790 100 390 5666979 2 3939356 6025716 3.74 2790 100 390 8482332 4 8177688 8482332 3.55 2790 100 390 9105481 5 8894881 9481360 3.53 2790 100 390 23810485 5 23705451 24357567 3.53 2820 100 420 23810485 5 23248880 25069184 4.33 157 Table 5.2 (continued ): Time (s) Intensity (µmol photons m-2s-1) Time since light change (s) Peak Position Chromosome 90% Confidence Interval logP lower upper 2820 100 420 5666979 2 3939356 6647145 3.79 2820 100 420 2902408 4 2803493 3924075 3.75 2820 100 420 23247571 5 23246477 23247571 3.72 2820 100 420 10302880 4 10043931 10977564 3.62 2820 100 420 22697296 5 21864028 23012412 3.58 2820 100 420 3344784 3 2903318 3578598 3.53 2850 100 450 23798428 5 23253768 24816019 4.10 2850 100 450 2907158 4 2803493 3924075 3.98 2850 100 450 10423538 1 9516375 11553534 3.87 2850 100 450 10302880 4 10042015 10777260 3.67 2850 100 450 8974265 1 8969354 9362052 3.61 2880 100 480 2907158 4 2695395 4933872 4.80 2880 100 480 1585202 4 1241726 1613368 3.65 2880 100 480 13217139 1 12646750 13217139 3.63 2880 100 480 1132454 4 1128719 1237935 3.61 2880 100 480 23798428 5 23396016 24357567 3.63 2880 100 480 11412672 1 11159782 11466186 3.58 2910 100 510 2907158 4 2588286 5291404 5.72 2910 100 510 1585202 4 1238602 2441130 4.02 2910 100 510 23798428 5 23400832 24357567 3.69 2910 100 510 1132454 4 1128719 1132454 3.57 2940 100 540 2907158 4 2695395 4803015 4.37 2940 100 540 13217139 1 12646750 13217139 3.58 2940 100 540 11003558 4 11001770 11470264 3.58 2940 100 540 10777260 4 10613944 10977564 3.55 2940 100 540 23798428 5 23705451 24357567 3.54 2940 100 540 11412672 1 11159782 11412672 3.46 3000 100 600 2907158 4 2803493 3924075 3.78 158 Chapter 6 Conclusions and future directions Geoffry A. Da vis 159 6.1 Abstract. The work in this chapter is to contextualize the concepts presented within this dissertation as a whole as well as in the larger field of photosynthesis research. Questions that have emerged from the work presented in this dissertation and how they can be projected forward to better understand how photosynthesis occurs within cells in natural environments are discussed. 6.2 Conclusions. Advances in crop yield over the last century have not occurred simultaneously with increases in photosynthetic capacity (see Chapter 5). While this is not in itself a problem, the discrepancies between theoretically maximal photosynthetic yields of the light reactions versus the actualized potentials of crop species provides opportunities to nudge photosynthesis closer to optimum potentials. While the core electron transfer processes of the photosynthetic light reactions are well characterized from atomistic to mechanist levels, many of the regulatory factors that act on the core electron and proton transfer complexes are p oorly understood in their cellular and physiological context. This is due to a number of factors, including, but not limited to, long growth cycles of certain species, technical challenges of certain measurements, as well as a small (but expanding) reperto ire of plant model organisms. To overcome some of these challenges, high -throughput photosynthetic phenotyping instruments have been developed by multiple groups to gain mechanistic insight from large datasets of responses to stress (see Chapters 2 and 5 ). Expanding the methodologies used to measure photosynthetic parameters, combined with the increasing amount of genetic information available for photosynthetic organisms, can lead to improvements in our understanding of what processes limit photosynthesi s under natural conditions, how those processes are regulated, and if there are ways that it could be improved to minimize those limitations. 160 6.3 Future directions. The large impact that !! imposes on photosystem (PSII) electron transfer reactions (see Chapters 2, 3, 4) leads us to believe that photosynthetic organisms have evolved mechanisms to minimize the photodamage potential of !! and increases in recombination. The core electron transfer proteins of PSII are highly conserved across oxygenic photosynt hetic organisms (219), as well as being mechanistically and structurally conserved between PSII and anoxygenic reaction centers (220). However, unlike eukaryotic photosynthetic organisms, which usually posses a single copy of the psbA gene to produce D1, c yanobacteria often possess multiple psbA genes (219). A potential advantage of maintaining multiple psbA genes is the ability to tune PSII activity for environmental conditions. As such, specific mutations in D1 have been associated with alteration in the D1 pheophytin hydrogen bonding network, leading to alterations in PSII recombination and photodamage (145, 184). Whether these canonical Òhigh lightÓ psbA isoforms are adaptive advantages to minimize PSII recombination in fluctuating light environments cou ld provide one avenue to minimize light fluctuation induced PSII photodamage. Unlike cyanobacteria, higher plant chloroplast genomes contain a single psbA gene, preventing any direct modulation of the free -energy gaps via protein subunit replacement. However, the rapid and consistent increases in PSII recombination during fluctuating light (Chapter 4) leads to the proposal that organisms have either evolved mechanisms to minimize the severity of rapid !! increases, and/or that mechanisms have evolved to rapidly respond to the oxidative stress that occurs after !! mediated increases in reactive oxygen species. While it appears that regulating the redox state of Q A via nonphotochemical quenching (NPQ) is one mechanism to minimize the duration of increased PSII recombination yields (Chapter 4), other mechanisms may exist to minimize photodamage from fluctuating light (Chapter 3). 161 Identification of QTL that appear in a time -dependent manner (Chapter 5) may represent alleles responsible for mediating immediate versus late responses to fluctuating light, which could be tested via allele swapping between ecotypes via CRISPR/Cas9 targeted gene replacement. To more specifically identify mechanisms related to PSII recombination, delayed fluorescence could be measure d in the Arabidopsis thaliana multi -parent advanced generation inter -cross (MAGIC) population under fluctuating light. Similarly to the prompt chlorophyll fluorescence analysis (Chapter 5), a camera with a higher sensitivity could be used to measure change s in the yield of delayed fluorescence following fluctuating light treatments to map QTL responsible for differences in PSII recombination yield as well as differences in how rapidly the increased PSII recombination is relieved. This would be complementary to the analysis already performed, as overlapping QTL could provide better mechanistic insight for the causative allele. The sensitivity of NPQ mutants to fluctuating light (Chapter 4) suggests that regulation of steady -state NPQ quenching is importan t to minimize the levels of Q A- in the event of an immediate increase in light intensity. If Q A- is minimized, potential photosynthetic productivity is limited, however, the plant may remain better protected during light fluctuations. The cost benefit tradeoffs for plant yield in rapidly regulating Q A- via NPQ could also be analyzed in the MAGIC popu lation. Under short term fluctuations, variations in the kinetics and extent of NPQ existed within the MAGIC founding accessions as well as the mapping population. Long -term fluctuating growth under dynamic, fluctuating light utilizing chlorophyll fluoresc ence imaging during vegetative growth would allow both QTL analysis of photosynthetic and yield parameters over developmental stages, but also provide a large, genetically diverse but naturally occurring dataset of evolutionarily tuned photosynthetic respo nses to fluctuating growth. This would 162 provide information to suggest whether protection from !!-induced photodamage or rapid relaxation of NPQ provide greater yield benefits under natural light conditions. 163 APPENDIX 164 A.1 Descripti on of DeltaPsi.py code for simulating the light reactions of photosynthesis. 3 DeltaPsi.py is a program for first -order, exploratory simulations of the effects of capacitance, proton buffering capacity and counter -ion movements on the thylakoid proton moti ve force ( pmf ), trans -thylakoid electric field ( !!), stroma -lumen pH difference ( !pH), linear electron flow (LEF), the ratio of ATP/NADPH produced by LEF, the activation and recovery of the lumen pH -dependent form of non photochemical quenching (NPQ), termed q E, photosystem II (PSII) activity and recombination rates and 1O2 production. The code is based on the simulations in (2) because simulations using this code were validated by the review process in both the original publication and in a separate, follow up publications (146). The following describes changes in this version. 1) The code was updated to run on modern, open -source, cross -platform and freely -available Python platform. Examples below were performed using the open source Jupyter notebook platform (www.jupyter.org), but code can also be run on any Python platform. 2) To provide the maximal transparency and allow others to repeat and modify our code and simulations, the full code is made available on GitHub (www.github.com/protonzilla/Delta_Psi_Py ) with extensive annotation. Readers are encouraged to fork the code and add their modifications, improvements and tests. It is hoped that these open source tools will provide the capacity to extend or modify and extend and validate the simulations, especially to test new hypotheses. 3 This section originally published as electronic supplemental material for: Davis, GA, Rutherford, AW, & Kramer, DM. 2017 Hacking the thylakoid proton motive force for improved photosynthesis: modulating i on flux rates that control proton motive force partitioning into !! and !pH. Philos Trans R Soc Lond B Biol Sci, 372(1730). doi: 10.1098/rstb.2016.0381 165 3) A more efficient ODE solver, odeint ( scipy.integrate.odeint : https://docs.scipy.org/doc/scipy -0.18.1/reference/generated/scipy.integrate.odeint.html ), was used to increase the speed and accuracy of the simulations. 3) Several improvements in the formulae were made to include several recent adva nces, as outlined in the following. a) Explicit simulation of Q A and plastoquinone (PQ) redox states were included to simulate PSII quantum yields and to account for the effects of !! on PSI I recombination reactions. b) The lumen pH -dependence of the xanthophyll cycle and the protonation of PsbS were updated to simulate the pH dependencies reported in Takizawa et al. (47) and Zaks et al. (146). c) The effects of lumen pH on plastoquinol (PQH 2) oxidation at the cytochro me b6f complex were included to account for the observed Òphotosynthetic controlÓ of electron transfer by the lumen pH, using the model and p Ka values reported in (47). d) The driving force for ATP synthase reaction (and thus the efflux of protons) is now taken as being equal to the difference in the proton motive force ( pmf ) and the free energy storage in ATP ( !GATP ) divided by n, the stoichiometry of H + translocated through the ATP synthase over the number of molecules of ATP formed, as described in more detail below. e) A more realistic model for the kinetics of the cytochrome b6f complex w as developed, to account for the thermodynamic and kinetic effects of pmf components on these reactions. In particular, the code now considers the b6f complex to be fully reversible, so that both !! and !pH alter the equilibrium constant. 166 f) An equation was derived and implemented to calculate the quantum efficiency of PSII based on NPQ and Q A redox state. A.1.2 Description of updated code. The follo wing describes the equations used in the simulations, focusing especially on those that extend the previous versions. Details of the calculations are embedded in the code and described in the code and the text. A.1.2.1 The pmf and its partitioning into !! and !pH . The theory and equations used to describe pmf components are essentially as described in earlier work by Cruz et al., (2) and Zaks et al. (146), with two updates. First, the new code allows for addition of antiporters or symporters, as described in the code for KEA3, which is proposed to act as a K +/H+ antiporter in the thylakoid membrane (28, 158): the KEA reaction looks like this: !!"#$% !!!!!"#$%& !!!!!!"#$%& !!!!!"#$% ! (Eq. 1) where !!"#$% !, !!"#$%& !, !!"#$% !, and !!"#$%& ! represent protons in the lumen and stroma and K + ions in the lumen and stroma, respectively. The reaction is electroneutral, so the forward reaction will depend on ![H+]and ![K+] as: !!"# !!!!!"# !!!!!"#$% !!!!!"#$% !!!!!!"#$%& !][!!!!"#$% !]) (Eq. 2) v_KEA = k_KEA*(Hlumen*Kstroma - Hstroma*Klumen) (Python) where !!"#$% ! (Hlumen) , !!"#$%& ! (Hstroma), !!"#$% ! (Klumen) , !!"#$%& ! (Kstroma) are the concentrations of free H + and K + ions in the lumen and stroma, respectively. Next, the following is used to calculate K + flux, which depends on the permeability of electrogenic K + channels and the KEA reaction. The permeability of K + through the K + channel 167 is perm_K. The K + flux through the electrogenic channel depends on both the K + concentration gradient and the electric field, i.e. !!!!, !!!!!!!"!!"# !"!!!"#$%& !!!"#$% !!!! (Eq. 3) K_deltaG=(.06*np.log10(Kstroma/Klumen) - Dy) (Python) where !!!"#$%& !!!!!! and Dy= !!, the transthylakoid electric field (stroma -lumen) in V. The instantaneous net change in lumen [ K+] per unit time is given by: !!!!"!!!"# !!!!!!!!!!!!"#$% !!!!"#$%& !! (Eq. 3) net_Klumen = perm_K * K_deltaG*(Klumen+Kstroma)/2 + v_KEA (Python) where net_Klumen = !!!!". In this simplified form, we assume that the flux through the K + channel is proportional to the driving force and the average concentration of K + in the stroma and lumen. A.1.2.2 PS II recombination reactions and singlet O 2 production. Earlier work (e.g. (221-223) shows that both !! and !pH should increase PSII recombination rates, and thus should increase 1O2 production. We derived an equation for estimating recombination rate in PSI I is described in Davis et al . (36) considering the effects of !pH and !! on energetics of electron sharing between redox intermediates: !!"#$%& !!!!!!!!!!"#$%& !"!!!!!!!!!!!"!!!!"!!!!"!!! (eq. 4 ) v_recomb = k_recomb*QAm*(10* *(fraction_Dy_effect *Dy/.06 + fraction_pH_effect*(7.0 -pHlumen)) (Python) where v recomb is the rate of recombination, [S 2QA-] is the concentration of PSII centers with reduced Q A and the oxygen evolving complex in the S 2 state, krecomb the intrinsic rate of recombination from S 2QA- in the absence of !! and !pH, ! Estab the stabilization free energy of 168 the charge separated state, expressed in eV (36), fD! (fraction_Dy_effect) and f DpH (fraction_pH_effect ) represents the fraction of quasi -stable S -states (that are able to recombine) and are sensitive to !! and !pH respectively (see text for more detail) . A.1.2.3 Normalizations of parameters. One (trivial) difference with our earlier model Cruz et al., (2) is that content and concentration parameters are normalized to PSII content rather than thylakoid surface area. This will make it easier to account for lumen volume changes. We use the general parameters from the literature, as reviewed in Cruz et al., (2), including the following. From Cruz et al., 2001 "...we estimated that there were approximately 2 ) 10-13 mol of PSI and PSII cm -2Ó of thylakoid membrane, or 6 X 10 10 PSII complexes cm -2. If all PSII centers were hit, on average there would be 6 x 10 10 cm-2 protons delivered into the lumen. 2 ) 10-13 mol of PSII cm -2 thylakoid membrane 6.02 x 1023 molecules per mole ) 10-13 moles of PSII cm -2 thylakoid membrane 1.2 x 1011 PSII centers per cm -2 thylakoid membrane 1.2 x 1011 protons cm -2, into 0.8 10 -9 L c m-2 i.e. 23 10 -6 moles/L are moved, equivalent to a change in the concentration of protons (both bound and unbound) of 2.3 10 -5 moles or about 12 µM. With 0.8 x 10 -9 L lumen volume cm -2 thylakoid membrane, yield 1.2x10 11 PSII centers per cm -2 thylakoid membrane, or 6.7x 10 -21 L per PSII center. Therefore, one turnover of PSII should introduce about 1.7 10 -24 moles H+ /6.7 10 -21 L equivalent to a change in concentration of about 2.5 10 -4 M. Given the large lumen buffering capacity, % =0.03 M/pH unit (22), the vast majority of these protons become buffered, and a single turnover flash should yield a !pH change of only 169 about 0.008 units. On the other hand, the capacitance of the thylakoid membrane is expected to be small, about 0.6 µF/cm2 (see tex t) so that moving one charge per PSII center should produce a substantial !!, Hitting all PSII centers with a single turnover flash should move 1.2 x 10 11 charges cm -2 or 1.2 x 1011 charges/ 6.242)1018 charges coulomb -1 =1.9E-8 C, and with 1.9E-8C/0.6 E-6C/V , and thus: !!(flash)=0.033 V We use this value to indicate the effective !! for a transthylakoid movement of one charge per PSII equivalent. A1.2.4 The reactions of the cytochrome b6f complex. The overall reaction for the cytochrome b6f complex is takes to be: PQH2 + PC(ox) ! b6f ! PQ + pmf + PC(red) (Eq. 6) where PQ and PQH 2 are the oxidized and reduced forms of plastoquinone, PC(ox) and PC(red) are the oxidized and reduced forms of plastocyanin, and pmf is the proton motive force. Ther e are several factors to consider. First, there is a kinetic effect of lumen pH on the binding of PQH 2 to the Rieske Fe 2S2 protein (224), with a pK a near 6 -6.5 (see (47) and references within), so that: b6f(H +) ! b6f(active), pKreg~6.5 (Eq. 7) where b 6f(active) is the deprotonated, active and b 6f(H +) is inactive forms of the complex. This is a kinetic constraint, likely related to the deprotonation of a His residue on the Rieske FeS protein , see Crofts and Wang (224) for a detailed description of the ex perimental bases of this interpretation in the cytochrome bc1 complexes. The term, pKreg, describes the pH -dependence 170 of activation of the cytochrome b6f complex. The empirically measured value is between 6.0 and 6.5 (47). Next, we need to consider the redox states of PQH 2, PC as well as the !! and !pH components of pmf . Because the Q -cycle releases 2 H + into the lumen for each electron passed from PQH 2 to PC, there should be a thermodynamic constraint to the forward reaction: 0.5PQH2 + b 6f(active) + PC(ox) --k_b6f ! PQ + b6f(H +) + PC(red) + 2 H+(lumen) (Eq. 8) This is somewhat simplified in that the intermediate reactions involving electron transfer through the low potential cytochrome b chain are ignored and it is assumed that the electron from the semiquinone generated in the Q o site of the complex is immediately transferred to PQ, so that while one PQ is produced during the Q o site reaction, it is partially reduced with the net effect of oxidation of a 0.5 PQH2. The forward rate constant is k_b6f, but the reaction is reversible, so that : 0.5PQH2 + b 6f(active) + PC(ox) !!k_b6f_reverse -- PQ + b6f(H +) + PC(red) + 2 Hin (Eq. 9) k_b6f_reverse is a function of pmf because the Q -cycle in the forward direction works against both !pH and !!. Note that this thermodynamic effect is in addition to the kinetic effect on the deprotonation of the Rieske protein. We simulate this as follows: !!"!!!!!!!!"!"!"!"#!!!!"!"#!!!!!"# (Eq. 10) Keq_b6f = E m(Pc(ox)/PC(red)) - Em(PQ/PQH 2) Ð n*pmf (Python) 171 where !!"!!!! is the equilibrium constant for the forward reaction, and E m(Pc(ox)/PC(red))= ) = 0.370 V and E m(PQ/PQH 2)=0.11 V at pH=7, are the effective redox midpoint potentials for the PC and PQ couples at pH=7, but are pH -dependent so th at: Em(PQ/PQH 2) = 0.11 V - (7-pHlumen ) * 0.06 (Eq. 12) In other words, the overall equilibrium constant is determined by the redox potentials of the donor and acceptor together and the pmf . We use unity as the scaling factor for the pmf contributions beca use one proton translocated to the lumen per e - transferred (together with one e- charge moved from the p - to the n -side) equilibrium. !!!!!!"#"!$" !!!!!!!"#$%#& !!"!!!! (Eq.13) k_b6f_reverse = k_b6f / Keq_b6f (Python) In principle, we could simulate the effects of changing PQH 2 and PC redox states in two ways, either using the simulated concentrations of PQH 2 and PC together with the standard E '0 (Em) values, or accounting for the concentrations in the E m values. We chose the former because it better fits the form of the ODE equations and is a bit simpler to calculate. Thus, !!!!!!"#!!"!"!!!!!!"#$%#& !!"!!"!"#!!!!!!!!"#"!$" (eq. 14) v_b6f=[PQH2] [PC_ox] k_b6f - [PQ] [PC_red] k_b6f_reverse (P ython) The midpoint potential of PC is pH -independent under our conditions, but E'0(PQ/PQH2) = 0.11 V at pH=7, but pH -dependent (see Eq. 12) so that: 172 !!"!!!!!!!!"!"!"!"#!!!!"!"#!!!!!"# !!!!!"#!!!!!!!!!!!!"!!!!!"#$%&' !!!!!!!!!!!!!"# (Eq. 15) Keq_b6f = E'0(Pc_ox/PC_red) - E'0(PQ/PQH2) - pmf = 0.370 - 0.11 + .06 * (pHlumen - 7.0) Ð pmf (Python) So, the full set of equations in Python is: Em7_PC=0.37 Em_PC=Em7_PC Em7_PQH2 = 0.11 Em_PQH2= Em7_PQH2 + 0.06*(pHlumen -7.0) #(Eq. 16) Keq_b6f = 10**((Em_PC - Em_PQH2 - pmf)/.06) #(Eq. 17) k_b6f_reverse = k_b6f / Keq # (Eq. 18) v_b6f=PQH2 PC_ox k_b6f - PQPC_red k_b6f_reverse # (Eq. 19) A.1.2.5 Calculating Phi2. The following is a derivation for determining Phi2 from NPQ and Q A redox state . The equations and derivations are based on those presented in Kramer et al. (100). Recall that NPQ is the ratio of: !"#!!!"# !!!!! (Eq. 20) where k NPQ , kf and k d are the intrinsic rate constants for NPQ, fluorescence and non -radiative decay of excitons in the photosynthetic antenna. Also, maximal PSII quantum yield is: !!!!!"# !!!"!!!!!!!!"!!!!! (Eq. 21) 173 !!!!!!"# !!!!!!!!!"!!"!!!!!!!!!!!" (Eq. 22) !!"!!!!!!!!!!!!!"# !!!!!!! (Eq. 23) where k pc is the maximal rate constant for PSII photochemistry. The realized !!! at any time is given by: !!!!!!!!!!"!!!!!!!!"# !!!!!!!" (Eq. 24) where [Q A] is the fraction of open PSII centers with oxidized Q A. !!!!!!!!!!!!!"# !!!!!"!!!!"!!!!!!!!!!!"# !!!!"!!!!!!!!!!!!"!!!"# !!!!"!!!!!!!!!!!!"!!"# !!!!!!!!!" (Eq. 25) !!!!!!!!!!!!!!!!"!!"# !!!!!!!"!!!!!!!!"!!!"#!!!!!"# !!!!! (Eq. 26) !!!!!!!!!!!"# !!!!!!! (Eq. 27) !!!!!!!!"# !!!!!!!! (Eq. 28) !!!!!!!!"# !!!!!!!! (Eq. 29) In Python, this translates to: def Calc_Phi2(QA, NPQ): Phi2=1/(1+(1+NPQ)/(4.88*QA)) return Phi2 174 REFERENCES 175 REFER ENCES 1. 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