MIAMI UNIVERSITY the Graduate School

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Isha Kalra

Candidate for the Degree

Doctor of Philosophy

______Rachael Morgan-Kiss, Director

______Xin Wang, Reader

______Donald J. Ferguson, Reader

______Annette Bollmann, Reader

______Carole Dabney-Smith, Graduate School Representative

ABSTRACT

ROLE OF CYCLIC ELECTRON FLOW (CEF) AND PHOTOSYSTEM I (PSI) SUPERCOMPLEX FORMATION DURING ACCLIMATION TO LONG-TERM SALINITY STRESS IN GREEN ALGAE: A COMPARATIVE STUDY

Isha Kalra

Photosynthesis is one of the most important processes on Earth by which organisms convert solar energy into usable forms of energy. Linear electron flow (LEF) and cyclic electron flow (CEF) constitute two major pathways in photosynthesis. While LEF leads to production of both ATP and NADPH, CEF only produces ATP that helps balance the ATP:NADPH ratio required for carbon fixation. CEF also plays a major role during acclimation to several environmental stressors. However, the regulation and mechanism by which CEF operates is still not clearly understood. Recent studies have shown that formation of a protein supercomplex with PSI appears to be essential for induction of CEF in several model organisms. However, both supercomplex formation and CEF induction have been mainly studied under short-term, transitory stress conditions. In addition, the role and mechanism by which organisms may rely on CEF to survive in their natural habitat and acclimate to stress over a long period of time has not been considered. In this study we compared how three photosynthetic organisms (one model alga, Chlamydomonas reinhardtii; two extremophiles, C sp. UWO241 and C. sp. ICE-MDV) utilize CEF to cope with their natural environment and adapt to steady-state environmental stress. To that end, the objectives of this thesis were i) to elucidate the role of CEF in long-term salinity acclimation ii) to understand the downstream changes associated with increased CEF, and iii) to identify whether PSI-supercomplexes are associated with increased CEF during salinity acclimation. We hypothesized that a stable PSI-supercomplex is required for high CEF, which in turn supports strong carbon fixation capacity for production of downstream metabolic products important for long-term acclimation to salinity stress. We showed for the first time, that increased CEF in UWO241 leads to excess ATP production and rewiring of downstream metabolism under high salinity. Next, we showed that a laboratory evolved salinity-tolerant strain of model C. reinhardtii uses constitutive upregulation of CEF to deal with salinity stress, which is in-turn associated with increased non-photochemical quenching and rewired carbon metabolism. Last, we show that CEF is involved in salinity acclimation in all three Chlamydomonas species, regardless of their salinity tolerance. We also show that PSI-supercomplexes are associated with increased CEF in these species. Characterization of high-salt supercomplex of C. reinhardtii revealed that it shares many similarities with the extensively described state 2 supercomplex, and that supercomplex composition might be species dependent rather than stress dependent.

ROLE OF CYCLIC ELECTRON FLOW (CEF) AND PHOTOSYSTEM I (PSI) SUPERCOMPLEX FORMATION DURING ACCLIMATION TO LONG-TERM SALINITY STRESS IN GREEN ALGAE: A COMPARATIVE STUDY

A DISSERTATION

Presented to the Faculty of

Miami University in partial

fulfillment of the requirements

for the degree of

Doctor of Philosophy

Department of Microbiology

Isha Kalra

The Graduate School Miami University Oxford, Ohio

2021

Dissertation Director: Rachael Morgan-Kiss

Isha Kalra

2021

TABLE OF CONTENTS

LIST OF TABLES VI LIST OF FIGURES VII DEDICATION VIII ACKNOWLEDGEMENTS IX CHAPTER I. INTRODUCTION 2 1.1 INTRODUCTION 2 1.2 PHOTOSTASIS AND ENVIRONMENTAL STRESS RESPONSE 3 1.3 CYCLIC ELECTRON FLOW 5 1.3.1 PSI-Supercomplex formation 6 1.4 EXTREMOPHILES 8 1.4.1 Psychrophilic phototrophs of McMurdo Dry Valleys, Antarctica 8 1.4.1.1 The model for psychrophilic photosynthesis: Chlamydomonas sp. UWO241 9 1.4.1.2 New phototroph from Lake Bonney: Chlamydomonas sp. ICE-MDV 10 1.5 OBJECTIVES OF THE THESIS 11 1.6 REFERENCES 16

CHAPTER II. CHLAMYDOMONAS SP. UWO241 EXHIBITS CONSTITUTIVELY HIGH CYCLIC ELECTRON FLOW AND REWIRED METABOLISM UNDER HIGH SALINITY. 34 2.1 INTRODUCTION 34 2.2 MATERIALS AND METHODS 36 2.2.1 Culture conditions, growth physiology. 36 2.2.2 Low temperature (77K) fluorescence spectra. 36 2.2.3 P700 oxidation-reduction and cyclic electron flow. 37 2.2.4 In vivo spectroscopy measurements. 37 2.2.6 Thylakoid isolation. 38 2.2.7 SDS-PAGE and Immunoblotting. 38 2.2.8 Supercomplex isolation. 38 2.2.9 Sample preparation for proteomics. 39 2.2.10 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS) 40 2.2.11 Gas Chromatography - Mass Spectrometry. 41 2.3 RESULTS 42 2.3.1 UWO241 is adapted to low temperature and high salt. 42

iii 2.3.2 UWO241 possesses constitutively high rates of CEF. 42 2.3.3 Isolation of a PSI-supercomplex in UWO241. 43 2.3.4 Protein composition of the supercomplex. 44 2.3.5 Whole cell proteome analysis. 45 2.3.6. Primary metabolome analysis. 46 2.4 DISCUSSION 47 2.5 REFERENCES 64 2.6 APPENDIX 74

CHAPTER III. COORDINATED RESPONSE OF CYCLIC ELECTRON FLOW, PHOTORESPIRATION AND TRANSIENT STARCH SYNTHESIS IN A HIGH SALT-EVOLVED STRAIN OF CHLAMYDOMONAS REINHARDTII 102 3.1 INTRODUCTION 102 3.2 METHODS 106 3.2.1 Growth conditions and evolution of high salt evolved strain of C. reinhardtii 106 3.2.2 Experimental set-up 106 3.2.3 PSII measurements and oxygen evolution rates 107 3.2.4 P700 photo-oxidation and cyclic electron flow 107 3.2.5 Protein extraction and sample preparation for proteomics 108 3.2.6 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS) 109 3.3 RESULTS 109 3.3.1 The high salinity evolved strain has faster growth rate 109 3.3.2 The evolved strain maintains high photosynthetic capacity and constitutive upregulated NPQ under low and high salinity 110 3.3.3 The evolved strain has higher oxygen evolution and respiration rates compared to the parent strain 111 3.3.4 The evolved strain displays constitutive high rates of PSI-CEF 111 3.3.5 Proteomic comparison of the evolved and the wild type strains under high salinity 112 3.3.5.1 Sub-cellular localization 112 3.3.5.2 Gene ontology 113 3.3.5.2 Kegg Orthology Biological Pathway 113 3.4 DISCUSSION 118 3.5 REFERENCES 153

3.6 APPENDIX 167

CHAPTER IV. IMPACT OF SALINITY-TOLERANCE VERSUS-ACCLIMATION ON THE STRUCTURE AND FUNCTION OF THE PHOTOCHEMICAL APPARATUS: A COMPARATIVE STUDY 170 4.1 INTRODUCTION 170 4.2 METHODS: 174 4.2.1 Culture conditions, growth physiology. 174 4.2.2 State transition induction 174 4.2.3 Low temperature (77K) fluorescence spectra. 175 4.2.4 PSII fluorescence state transition measurement 175 4.2.5 SDS-PAGE and Immunoblotting. 175 4.2.6 P700 oxidation-reduction kinetics 176 4.2.7 Supercomplex isolation. 176 4.2.8 Sample preparation for proteomics. 177 4.2.9 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS) 177 4.3 RESULTS 178 4.3.1 Salinity tolerance the three Chlamydomonas species 178 4.3.2 Photosystem I activity 179 4.3.3 Effect of long-term stress high salinity acclimation on short-term state transition response 180 4.3.4 Thylakoid protein phosphorylation 182 4.3.5 Assembly of protein supercomplexes under high salinity 183 4.3.6 Proteome analysis of high salinity-associated supercomplexes 185 4.4 DISCUSSION 186 4.5 REFERENCES 199 4.6 APPENDIX 213

CHAPTER V. CONCLUSION 216

LIST OF TABLES

2.1 Components of UWO241 supercomplex under high salinity 53 3.1 Growth characteristics of Wildtype and Evolved strain under low and high salinity 136 3.2 Significantly regulated proteins of the Wildtype strain under high salinity 137 3.3 Significantly regulated proteins of the Evolved strain under high salinity 144 4.1 Major proteins involved in high salt and state 2 supercomplexes of C. reinhardtii 198

LIST OF FIGURES

1.1 Two modes of photosynthetic electron transport 14 1.2 Differences between stress, acclimation and adaptation 15 1.3 Model of East Lake Bonney 16 2.1 P700 re-reduction kinetics of UWO241 54 2.2 Photosynthetic properties of UWO241 using Electrochomic shift 55 2.3 Supercomplex formation in UWO241 vs C. reinhardtii 56 2.4 77K fluorescence spectra of protein complex of UWO241 and C. reinhardtii 57 2.5 Heatmap of proteins of UWO241 under low and high salinity 59 2.6 Changes in photosynthesis and metabolism of UWO241 under high salinity 61 2.7 Model of photosynthesis and metabolism of UWO241 under high salinity 62 3.1 Comparison of growth of different C. reinhardtii strains 125 3.2 PSII parameters of Wildtype and evolved strain under low and high salinity 126 3.3 Oxygen evolution and dark respiration rates of wildtype and evolved strain 127 3.4 P700 kinetics of wildtype and evolved strain of C. reinhardtii 128 3.5 Heatmap of proteins of Wildtype and evolved strain under high salinity 129 3.6 Analysis of significantly regulated proteins of Wildtype and Evolved strain 130 3.7 Categorization of differentially regulated proteins into gene ontology 131 3.8 Categorization of differentially regulated proteins into Kegg Orthology 132 3.9 Analysis of common proteins of Wt and Ev strain 133 3.10 Analysis of unique proteins of Wt and Ev strain 135 4.1 Growth curve under salinity gradient for the three Chlamydomonas species 192 4.2 P700 oxidation/reduction kinetics of the three Chlamydomonas species 193 4.3 State transition tests after acclimation to low and high salinity 194 4.4 Thylakoid phosphorylation pattern of the three Chlamydomonas species 195 4.5 Isolation of supercomplexes from conditions promoting CEF in Chlamydomonas species 196 4.6 Proteome comparison of supercomplex fractions from state 2 and high salt cultures 197 5.1 Changes in photosynthesis and metabolism after acclimation and adaptation 220

vii

DEDICATION

This thesis is dedicated to my younger brother and only sibling, Akshay, who was the light and the joy of my life. They say a mother’s love is unmatched, but I believe a sibling’s love is boundless. Kind and beautiful, there is no one who has showered me with more love and joy than him. He was my motivation to pursue research and my support all throughout my life. He will continue to motivate me in my journey beyond and I hope to keep honoring his memory in the future.

viii

ACKNOWLEDGEMENTS

The last several years of my PhD have been an intense learning experience and there are several people who have helped me through that journey. First and foremost, I would like to thank my PhD supervisor, Dr. Rachael Morgan-Kiss, for her absolute support and guidance all throughout my PhD. I learned how hard-work and joy in your work can go hand-in-hand from her. I am also thankful for her for cultivating a great collaborative environment to pursue research and for always willing to think of creative ways to get work done. A huge thanks to Rachael for providing amazing opportunities and for offering to help in times of need. The last semester was especially tough, and I am extremely thankful for her patience and support during that time. Rachael created a home away from home for me, and I will always be grateful for that.

I would also like to thank all my committee members, who have helped me critically analyze my project and provide much valuable support and help. Thank you, Dr. Wang, for always motivating me to be my best self and offering your support during tough times. Thank you Dr. Bollmann and Dr. Ferguson for providing with me valuable feedback and interesting suggestions, and for your support. I would also like to thank Dr. Actis, for helping me learn the value of consistent hard-work and for always showing faith in me. A big thanks to Dr. Dabney- Smith for helping me fall in love with proteins and for being amazingly supportive all throughout.

My PhD journey would not be complete without the help, support and love of my friends. Thank you Shasten, Shrameeta, Mariah, Jananie and Jyoti for being my pillars of support and making my time at Miami fun filled. I would also like to thank Zev, Nathan, Chandiramani and Sabita for their friendship and kindness. Thank you Shaz for making the Antarctic field season so memorable and fun, I wouldn’t have been able to survive without you (Ishasten forever!). I had the most wonderful time with you and Zev as my lab-mates, couldn’t have asked for better ones. Thank you Shramy, for helping me during the toughest time in my life, I will forever be grateful for your care and presence in my life. Thank you Jyo for being my fellow quirky friend and sharing my Doja fandom, I will deeply cherish our time together at Miami. Thank you, Mariah, for being the kind and sweet friend you are. My time in Miami couldn’t have been full of color

ix without you and our Office jokes, I will sorely miss our time in room 50. Also, a huge thanks to Amy, Bev and Greg for keeping the department running and always helping with a smile.

Words are not enough to convey my thanks for my family. First and foremost, my sweetest baby brother Akki, who filled my life with so much love and happiness that it would last me all my life. He has supported, helped and cared for me more than anyone has in my life. His absence in my life is so huge that it almost feels like a presence. I carry you with me every second of everyday Akki, I can’t express my gratitude for your presence in my life. I hope you are watching me from above and I will strive to carry your infectious kindness and light with me. Thank you, dad, for inspiring me to be my best self and for being most loving father one can imagine. I am so lucky to have been blessed with a dad like you and I hope to keep making you proud. Thank you, mom, for being the sweetest and most supportive mother. I am here because of your unconditional love. I would also like to thank my new family, my mother-in-law for her huge support and kindness during tough times, my sister-in-law Vandu for making me smile at her silliest jokes and being the younger sibling that I can pamper and last my father-in-law for his wise words and support.

Last but not the least, a huge thanks to the love of my life, Adit, for sharing this journey with me. This PhD is as much yours as is mine. Your kindness, love and support have helped me tremendously during this time. You are my biggest cheerleader, and I wouldn’t be here without you. Thank you from the bottom of my heart.

CHAPTER 1

Introduction

Chapter I. Introduction

The food demands of the future population have been proposed to increase by 60% by 2050. Concurrently, climate change-associated declines in productivity of major food producing plants have also been observed: a trend which it is anticipated to accelerate in the near future (Alexandratos & Brunismas, 2012). To combat the increase in demand and overcome the environmental problems like drought, heat stress, high salinity brought by climate change, there is a need to engineer crops to survive and maintain or increase productivity in changing and extreme habitats (Kramer & Evans, 2011). Many organisms that survive in extreme habitats have evolved mechanisms that enable growth and maintenance of photosynthesis under numerous abiotic stresses (R. M. Morgan-Kiss, Priscu, Pocock, Gudynaite-Savitch, & Huner, 2006). To date, only a small handful of these photosynthetic extremophiles have been thoroughly described (Dolhi, Maxwell, & Morgan-Kiss, 2013). I hypothesize that by understanding mechanisms of photosynthetic adaptation to long-term, permanent environmental stress, we can discover novel strategies which can be translated to stress tolerance in crops. This project will address this hypothesis by focusing on one essential, poorly understood stress-acclimation pathway. Cyclic electron flow (CEF) is an essential pathway in photosynthesis that leads to ATP production as well photoprotection under short-term stress (Suorsa, 2015). The role of CEF in long-term stress acclimation in organisms living under permanent environmental stress is currently not fully understood. This project focuses on CEF as a strategy for long-term stress survival in two Antarctic psychrophilic green algae: a well-studied species, Chlamydomonas sp. UWO241 and a newly isolate, C. sp. ICE-MDV. We compare the physiology of these two extremophiles with that of the model alga, C. reinhardtii as well as a salt-tolerant evolved strain (Ev-HS) of the same organism.

1.1 INTRODUCTION

Photoautotrophic eukaryotes like green algae utilize oxygenic photosynthesis to capture abundant sunlight to fix carbon dioxide and produce oxygen as a byproduct. Chlorophytes (green algae) are the direct ancestors of modern land plants, and thus are useful single-celled models for their more complex counterparts. The thylakoid membrane in the chloroplast of plants and eukaryotic algae houses the photosynthetic electron transport chain (PETC). The PETC is comprised of multiple membrane-bound pigment-protein complexes which work collectively to

capture light energy, conduct electron charge separation, and ultimately produce ATP and NADPH (Roach & Krieger-Liszkay, 2014). Much of the PETC-derived ATP and NADPH are consumed by carbon fixation via the Calvin-Benson-Bassham (CBB) pathway (Dijkhuizen & Harder, 1985). The major light harvesting-energy transduction protein complexes flanking the PETC are photosystem II (PSII) and photosystem I (PSI). The antenna chlorophyll-pigment- protein complexes light harvesting complex II (LHCII) and I (LHCI) absorb the photons from light and transfer them to PSII and PSI respectively, where the excitation of chlorophyll releases electrons. Electrons are shuttled from PSII to PSI via the plastoquinone pool (PQ pool),

cytochrome b6f (cyt b6f) and plastocyanin (PC), generating NADPH and a proton gradient at cytb6f that drives ATP synthase to produce ATP. This basic process describes the classic “z- scheme” or linear electron flow (LEF) (Fig. 1.1) (Govindjee, Shevela, & Björn, 2017). Plants and algae also possess multiple alternative pathways which are utilized under different environmental scenarios to balance ATP and NADPH requirements. These alterative pathways will be discussed in the context of environmental stress response in more detail below.

1.2 PHOTOSTASIS AND ENVIRONMENTAL STRESS RESPONSE

Photosynthetic organisms must continuously adjust the PETC to keep the supply of ATP and NADPH closely matched with downstream metabolic energy needs (Cardona, Shao, & Nixon, 2018). Most significantly, there is a natural imbalance between the rates of fast, temperature- insensitive photochemical reactions through which light is absorbed and electrons are released in the PETC, and the downstream C, S and N metabolism which is much slower and temperature- sensitive (Kramer & Evans, 2011). Photosynthetic organisms have evolved to sense and avoid this imbalance through a number of processes which modify either PETC-derived energy production or metabolism-dependent consumption of ATP and NADPH. This maintenance state is called ‘photostasis’, where the amount of light energy absorbed and converted to chemical energy is relatively balanced with the amount needed to drive downstream metabolism (Hüner et

-1 al., 2012). Photostasis is defined using the equation sPSII.EK = t , where sPSII is the effective

PSII absorption cross-section, EK is the irradiance (I) at which the photochemistry is balanced with photosynthetic capacity and t-1 is the turnover rate for the metabolic sinks (Huner et al., 2003).

Many environmental stresses can exacerbate imbalances in the energy demands (R. M. Morgan-Kiss et al., 2006). These imbalances put the photosynthetic organisms in a state of high

-1 excitation pressure (HEP) which can be represented as sPSII.EK > t . As a result of HEP, the PQ pool and the PETC are in over-reduced redox state due to electron buildup in the photosynthetic machinery. This over-reduction of the plastoquinone pool will result in damage to the photosynthetic apparatus by formation of reactive oxygen species (ROS) via deposition of excess electrons on oxygen molecule. Consequently, the redox status of the PQ pool is a key sensing mechanism for HEP and photostasis (Hüner et al., 2012). As HEP is a result of energetic imbalances, any change in the environment of a photoautotrophic organism is reflected in the redox state of PQ pool. Thus, photosynthetic organisms kick-start acclimatory responses after sensing the redox state of the PQ pool and remodel the structure and function of their photosynthetic apparatus to return to a state of photostasis. In their natural environment, organisms must respond to various levels of environmental stress (including high light, low temperatures, high salinity, nutrient deficiency), which may last for only a few minutes (short-term or transient) or persist for days to years (long-term) (Fig. 1.2) (W. Huang, Zhang, & Cao, 2010; Lucker & Kramer, 2013; Strand et al., 2015; Yamori & Shikanai, 2016). During transient stress that leads to HEP, organisms respond by using short- term acclimatory mechanisms. State transitions are classic short-term mechanism by which organisms divert energy from PSII to PSI via reversible phosphorylation of major LHCII complexes, followed by migration of phospho-LHCII from PSII to PSI (Lemeille & Rochaix, 2010). LHCII attaches to PSI to reduce the amount of light absorbed at PSII and avoids photooxidative damage. State transitions and/or an over-reduced PQ pool are also associated with activation of alternative electron transport pathways. Another mechanism by which organisms acclimate in the short-term is through initiation of non-photochemical quenching (NPQ) response through activation of the xanthophyll cycle (Müller, Li, & Niyogi, 2001). During NPQ response organisms dissipate excess absorbed energy and reduce excitation pressure on the PETC, specifically to prevent PSII photodamage. In contrast with short-term acclimation, long-term stress organisms respond by global adjustments in the cell proteome, including changing protein levels of components of the PETC (Kono & Terashima, 2014). This can involve changes in the antenna-size of the photosystems, particularly major LHCII proteins to reduce/increase the amount of light absorbed (Melis, 1991).

Another typical response is alterations in PSII:PSI stoichiometry by changing levels of core proteins of one or both photosystems to balance the amount of energy absorbed by the two photosystems (Dietzel, Bräutigam, & Pfannschmidt, 2008). Moreover, some photosynthetic

organisms can also adjust CBB enzymes and intermediates, to increase rates of CO2 assimilation or photorespiration, acting as electron sinks, thereby relieving any HEP on the photosynthetic apparatus (Huner, Öquist, & Sarhan, 1998). To this end, green alga Dunaliella salina accumulates CBB enzymes to increase carbon fixation and photosynthetic capacity in response to high salinity stress (Liska, Shevchenko, Pick, & Katz, 2004).

1.3 CYCLIC ELECTRON FLOW

As mentioned above, the canonical route for PETC is the classic “z-scheme” which shuttles + electrons from H2O through PSII and PSI, ultimately depositing them onto NADP . This so- called linear electron flow (LEF) from PSII to PSI produces 2.57 ATP and 2 NADPH per two electrons. However ATP and NADPH production solely from LEF creates a conundrum, as the

downstream carbon fixation requires 3 ATP and 2 NADPH for assimilation of 1 CO2 molecule in the Calvin-Benson-Bassham cycle (Kramer & Evans, 2011). Thus, LEF alone does not provide ATP: NADPH in the correct ratio to sustain carbon fixation, and this does not even account for either additional needs of other metabolic pathways or stress-related changes in energy requirements. Plant and algae have evolved to deal with this apparent issue through activation of various alternative electron transport pathways which serve the purpose of titering PETC-derived energy flow (Roach & Krieger-Liszkay, 2014). One of the most active pathways,

cyclic electron flow (CEF), consists of electron transfer from PSI to plastoquinone to cyt b6f and back to PSI via plastocyanin (Fig. 1.1). As the electrons are cycling back to PSI, CEF only produces ATP via acidification of lumen by cyt b6f and no NADPH is produced (Yamori & Shikanai, 2016). Thus, CEF is generally accepted as an essential process in mitigating the imbalance between photosynthesis and carbon fixation. CEF is essential for producing additional ATP and also contributes significantly to photoprotection. CEF provides photoprotection in two ways: 1) triggering of NPQ by rapid acidification of lumen, thereby avoiding PSII damage, and 2) by reducing the PSI acceptor side limitation in condition of PSI photoinhibition (Finazzi & Johnson, 2016). The latter function is particularly significant: while PSII possesses a robust repair cycle, PSI does not (Gururani,

Venkatesh, & Tran, 2015). While the contribution of CEF to photosynthesis under non-stress conditions is <15%, the proportion of CEF relative to LEF increases drastically under conditions where the need for ATP and proton motif force (pmf) for photoprotection increases, a situation which is prevalent under many common environmental stresses, such as low temperature (W. Huang et al., 2010), drought (W. Huang et al., 2013), anoxia (Terashima et al., 2012), nutrient deficiency (Saroussi, Wittkopp, & Grossman, 2016). Moreover, environmental stress is often associated with HEP: CEF contributes to survival under stress conditions by reducing HEP in two major ways: (1) producing extra ATP, and (2) triggering dissipation of excess absorbed light through ΔpH-mediated NPQ (Yamori & Shikanai, 2016). While there are many unanswered questions regarding the regulation and mechanism of CEF, there are at least two main pathways for CEF: (1) the antimycin sensitive pathway using PGR5/PGRL1 (Proton Gradient Regulator 5/Proton Gradient Regulator Like protein, (2) the antimycin insensitive pathway utilizing NDH complex (NADPH dehydrogenase) for electron transfer (Suorsa, 2015). There is still a debate regarding the ubiquitous nature of these pathways, but it is recognized that the PGR5/PGRL1 pathway plays a larger role in green algae and the NDH pathway in plants (Yamori & Shikanai, 2016). Given the role of CEF in energy dissipation and ATP synthesis for stress acclimation, it makes an attractive target for manipulating plants for increased production in unfavorable conditions given the future food production demands and lack of agricultural land (Alexandratos & Brunismas, 2012). Studies to date have almost exclusively focused on the role of CEF under short-term stress in green algae (Iwai et al., 2010; Lucker & Kramer, 2013; Takahashi, Clowez, Wollman, Vallon, & Rappaport, 2013; Terashima et al., 2012); however, the role of CEF in long-term stress acclimation is not fully understood.

1.3.1 PSI-Supercomplex formation

Since CEF is a crucial and highly regulated process that needs to be flexible with the changing environment, the mechanism of its operation needs to be dynamic and operate over different time scales. While the CEF mechanism is not fully understood, formation of protein ‘PSI- supercomplexes’ appear to play an important role in CEF (Iwai et al., 2010). The assembly of these PSI-supercomplexes depends on the redox status of the PQ pool (Alric, 2015). Since their discovery, most studies have focused on isolating and characterizing supercomplexes under transient, short-term stress conditions where CEF is activated only on a short-term time scale.

The first stable supercomplex was isolated in 2010 by Iwai et al. in C.reinhardtii and was

composed of PSI, LHCII, cytochrome b6f, PGRL1 and Ferredoxin NADP Reductase (FNR) (Iwai et al., 2010). This supercomplex was isolated under conditions promoting state transition. State transition is a process that occurs within seconds to minutes in an organism after exposure to high excitation pressure, which leads to an over-reduced electron transport chain (Rochaix, 2007). In Chlamydomonas reinhardtii, the kinase STT7 senses the over-reduced PQ pool and phosphorylates LHCII. The phosphorylated LHCII then migrates towards PSI and forms a supercomplex to promote CEF to mitigate the HEP (Rochaix, 2007). In C.reinhardtii, another supercomplex was isolated under transient anoxia, an alternative procedure for inducing hyper-reduction of the PQ pool (Terashima et al., 2012). In the study the authors described the role of a calcium sensing protein (CAS) in the upregulation of CEF under anoxic conditions. Under the anaerobic condition they isolated a PSI-cyt b6f supercomplex that also consisted of ANR1 (anaerobic response 1 protein) protein along with CAS. This multi- protein complex was also shown to interact with PGR5-like (PGRL1) protein. In 2016, Takahashi et al showed that another protein PETO, which is a transmembrane photoprotein, is also involved in regulation of CEF in anoxic conditions in C.reinhardtii (Takahashi et al., 2016).

PETO was shown to co-fractionate with PSI, cyt b6f, FNR, PGRL1 and ANR1 as a supercomplex. The structure of C.reinhardtii supercomplex was recently elucidated under anaerobic conditions (Steinbeck et al., 2018). The authors used single molecule fluorescence

correlation spectroscopy to show physical association between PSI, LHCI and cyt b6f. The study also suggests that the dissociation of lhca2 and lhca9 subunits of the LHCI complex is important for docking of cyt b6f and formation of PSI-LHCI-cyt b6f supercomplex. Most of the studies focused on identifying proteins involved in supercomplex formation during upregulated CEF use treatments that are not physiologically relevant, such as, anoxia, chemical inhibitors to induce state transition, etc. There is also a dearth of studies focused on understanding the mechanism of CEF regulation under steady state conditions (Lucker & Kramer, 2013). Although these previous studies have identified proteins involved in supercomplex formation and CEF regulation during transient stress in green algae, it is still not fully understood whether CEF is important in long-term stress acclimation, if so, which pathway is utilized and whether a supercomplex is associated with it.

1.4 EXTREMOPHILES

Extremophilic photosynthetic organisms survive and photosynthesize under conditions of extreme stress. Non-model organisms that are adapted to very long-term stress lasting 100s- 1000s of years represent under-exploited reservoirs of novel adaptive mechanisms (Aguilera, Souza-Egipsy, & Amils, 2012). Unlike model organisms that are often studied for their short- term stress response, extremophiles can be used to understand acclimation strategies employed in lifeforms which survive under very long-term stress. By contrast, most of the model plants and algae which are generally utilized for stress studies possess minimal tolerance to extreme levels of environmental stress (Shetty, Gitau, & Maróti, 2019). For example, CEF is mostly studied in the model organism C.reinhardtii under transient stress conditions that are sometimes even physiologically irrelevant (Takahashi et al., 2016; Terashima et al., 2012). The supercomplex that initiates CEF under these conditions is also unstable and has been difficult to characterize because of the dynamic nature of the stress. On the other hand, our understanding of the role of CEF under long-term stress as well as steady state conditions is lacking. Using extremophilic organisms to probe the mechanism of CEF and supercomplex formation would provide insightful knowledge on the role of CEF in long-term stress acclimation. This information can help develop a stable and reliable mechanism that can be further used to engineer organisms capable of surviving in extreme habitats.

1.4.1 Psychrophilic phototrophs of McMurdo Dry Valleys, Antarctica

The vast continent of Antarctica is one the most extreme environments for life to survive: Antarctic lifeforms are almost entirely microbial. The McMurdo Dry Valleys are the largest ice- free area on the continent (Priscu, 1998). Despite the polar desert environment of the dry valleys, there exists numerous ice-covered lakes and ponds. These lakes are covered with a thick ice sheet throughout the year and only microbial life (bacteria, protists, fungi, algae) is sustained in the water column (Fritsen & Priscu, 1999). Photoautotrophic and mixotrophic microalgae are found at specific depths of the photic zones of the water column and are one of the major players in the food web dynamics (Lizottel & Prism, 1994). One of the most studied lakes in the McMurdo Dry Valleys is Lake Bonney. Lake Bonney is divided into two lobes, East Lobe Bonney (ELB) and West lobe Bonney (WLB) via a sill barrier (Priscu & Spigel, 1996). ELB is 40 m deep and is covered with 4 m of ice sheet (Fig. 1.3). The microbes in Lake Bonney also

undergo 6-8 months of complete darkness during winter. During the 4-6 months of summer the organisms grow under 24-hour daylight. Due to the thick ice sheet, the photosynthetic active radiation (PAR) is only 1% surface values and decreases sharply through the water column, with the deeper layers getting <<20 μmol photons m-2 s-1 of irradiance. The spectral range of the penetrating light is also narrow (350-450 nm) (Lizottel & Prism, 1994). As the lake is perennially covered with ice, there is little vertical mixing of the lake water. As a consequence, the lake is permanently stratified with oligotrophic shallow layers and mesotrophic deep layers (Priscu, Wolf, Takacs, & Fritsen, 1999). There is a steep chemocline at 15 m after which the salinity level in the lake increases drastically, making the deeper layers of the lake hypersaline (Priscu & Spigel, 1996). Last, the dry valley lakes are generally oligotrophic, and ELB is extremely phosphorous deficient.

1.4.1.1 The model for psychrophilic photosynthesis: Chlamydomonas sp. UWO241

The unicellular green alga Chlamydomonas sp. UWO241 was isolated from ELB in the McMurdo Dry Valleys in 1995 from the deep photic zone, 17 m depth (R. M. Morgan-Kiss et al., 2006) (Fig. 1.3). Extensive studies on this organism have demonstrated that it is adapted to the permanent low-temperature and high salinity environment of the deep layers of East lake Bonney. UWO241 is a psychrophilic halotolerant green alga: it requires temperatures <15 oC and can tolerate salinity levels of up to 1 M NaCl (Dolhi et al., 2013). As a consequence of adaptation to low temperature, high salinity and extreme shade, UWO241 has completely remodeled its photosynthetic apparatus. One of the major low temperature adaptions in UWO241 is the remodeling of its photosynthetic membrane to be enriched with unsaturated lipids to increase membrane fluidity (R. Morgan-Kiss, Ivanov, Williams, Khan, & Huner, 2002). Not only that, to deal with extreme low temperatures, UWO241 contains many ice-binding proteins (Raymond & Morgan-Kiss, 2013). The PSII of UWO241 is also associated with large LHCII antenna, most likely as an adaption to the shade environment in the deep layers of the lake. Moreover, UWO241 also maintains a high PSII:PSI ratio and a low chlorophyll a/b ratio (Morgan, Ivanov, Priscu, Maxwell, & Huner, 1998). One of the most unique adaptations of UWO241 to long-term environmental stress is downregulation of short-term acclimatory mechanisms. Early studies on UWO241 demonstrated that it has an absent PSI peak in 77K spectra and does not undergo classic state transition

9 response when exposed to short-term high light stress (R. M. Morgan-Kiss, Ivanov, & Huner, 2002). Not only that, but it was also shown that UWO241 lacks LHCII polypeptide phosphorylation (Szyszka, Ivanov, & Hüner, 2007), unlike other green algae (Rochaix, 2007). Instead of phosphorylating LHCII, UWO241 phosphorylates several high molecular mass polypeptides (R. M. Morgan-Kiss et al., 2002; Szyszka et al., 2007). On the other hand, to deal with long-term extreme stress, UWO241 displays constitutive high rates of PSI mediated CEF under steady-state condition (Szyszka et al., 2007). The identity of the high molecular weight phosphorylated polypeptides was investigated and it was demonstrated that UWO241 preferentially phosphorylates a 1 MDa supercomplex of PSI (Szyszka-Mroz, Pittock, Ivanov, Lajoie, & Hüner, 2015). The supercomplex was stable under high salinity and in presence of phosphatase inhibitor NaF. This supercomplex composed of PSI, cyt b6f, PGR5, LHCI, FtsH and PsbA. Szyszka et al. also showed that phosphorylation of FtsH and PsbA proteins is important for supercomplex formation and its stability. Remarkably the composition and phosphorylation status of this supercomplex of extremophile UWO241 is different from the complexes described under transient conditions such as state transitions and anoxia for model C.reinhardtii (Steinbeck et al., 2018; Takahashi et al., 2016; Terashima et al., 2012). In a surprising discovery, recently Szyzska et al. showed that UWO241 possesses a unique state transition capability with an altered protein phosphorylation profile as compared to the model alga C. reinhardtii. It was suggested that UWO241 balances redox pressure under short-term stress using energy spillover between PSII and PSI, which gives rise to the unique state transition phenomenon (Szyszka-Mroz et al., 2019).

1.4.1.2 New phototroph from Lake Bonney: Chlamydomonas sp. ICE-MDV

One obvious question that arose from the studies in UWO241 is whether the novel adaptive strategies in this organism are commonplace among psychrophilic algae? To answer this question, a second photopsychrophile was recently isolated from the upper water column of ELB. This new isolated was named Chlamydomonas sp. ICE-MDV (ICE-MDV) because its 18S rRNA is 100% identical to that of an Antarctic marine species, C. sp. ICE-L (Raymond & Morgan-Kiss, 2013; Szyszka-Mroz et al., 2019). Recent studies have shown that contrary to previous assumptions, ICE-MDV dominates the chlorophyte populations, while UWO241 is very low in abundance (Dolhi et al., 2013; Dolhi, Teufel, Kong, & Morgan-Kiss, 2015). Unlike

UWO241, this organism was isolated from higher in the water column, which is characterized by high PAR levels, lower salinity, as well as lower nutrient levels. ICE-MDV also receives significantly higher PAR (~50 μmol photons m-2 s-1) as compared to shade-adapted UWO241 (Fig. 1.3). Although ICE-MDV is a psychrophile, it is unknown whether it can tolerate high salinity like UWO241. It is yet to be investigated whether ICE-MDV shows similar mechanisms to adapt to its extreme environment like state transition response, CEF rates and supercomplex formation as described in UWO241.

1.5 OBJECTIVES OF THE THESIS

Numerous studies have shown the benefit of CEF for energy balance and acclimation to environmental stress. Formation of a PSI-supercomplex appears to be essential for induction of CEF in several model plants and algae. However, both supercomplex formation and CEF induction have been mainly studied under short-term, transitory conditions. This leaves an obvious gap in the knowledge regarding whether these processes play roles in survival to stress under longer time periods. Moreover, several studies have focused on isolating supercomplexes required for CEF under transient or short-term stress; however, under such conditions the supercomplexes may be unstable so a precise mechanism for CEF has not been fully elucidated. The role and mechanism by which organisms may rely on CEF to survive their natural habitat and acclimate to stress over a long period of time has not been considered. There is a critical need to study CEF in relation to long-term stress acclimation and identify the components of supercomplex initiating CEF under these conditions. Unless we understand the role of CEF and its associated supercomplex during long-term and short-term stress, the exact mechanism and regulation of CEF will evade us.

In this study I will use non-model organisms (UWO241 and ICE-MDV) to understand the role of CEF in long-term stress acclimation. I will also investigate the role of supercomplex formation under steady-state physiologically relevant conditions of Lake Bonney habitat in both extremophiles. Extensive studies on UWO241 have identified many changes in the photosynthetic apparatus associated with its adaptation to permanent low temperature and high salinity; however, the downstream metabolic changes associated with such adaptation have not been studied. In this project I will also investigate the metabolic changes associated with high

11 salinity adaption in UWO241. Most CEF and supercomplex studies on C. reinhardtii have been conducted under transient stress conditions. In this project, I will investigate the role of CEF in the long-term acclimatory response of a laboratory salinity-evolved strain of C. reinhardtii. The overall objective of this project is to understand the mechanism of CEF and supercomplex formation, and their role, in long-term acclimation to environmental stresses associated with high salinity. Our central hypothesis is that a stable PSI-supercomplex supports sustained high CEF for downstream metabolic products important for long-term acclimation to salinity stress. The following questions and hypotheses will test the central hypothesis of my thesis:

1. Past studies have described ways in which UWO241 has modified its photosynthetic apparatus to deal with the permanent stress. Sustained CEF in UWO241 should alter the energy products (ie. ATP: NADPH) from photosynthesis. What are the consequences of sustained CEF on downstream metabolism in UWO241? I hypothesize that increased CEF in UWO241 supports a rewired metabolism facilitating upregulation of energy consuming metabolic products.

2. UWO241 is a model to study long-term stress adaptation and mechanism of CEF that operate in such conditions. As this organism cannot be genetically manipulated, it is not possible to make mutations in the proteins involved in the supercomplex to identify their essentiality. To overcome this problem, we propose to generate an evolved strain of the model green alga C. reinhardtii that survives higher salinity relative to the progenitor strain. I hypothesize that this evolved strain will exhibit a phenotype that is comparable with that of UWO241 with increased rates of CEF to acclimate under long-term salinity stress.

3. The Antarctic photopsychrophile Chlamydomonas sp. UWO241 has sustained high rates of CEF associated with a PSI-supercomplex, when exposed to long-term salinity stress. Another unique phenotype associated with the organism’s adaptation to environmental stress is the downregulation of short-term stress responses such as state-transitions. What is the role of CEF in acclimation to long-term stress and how does it affect short-term

12 stress response? I hypothesize that photosynthetic organisms acclimate to long-term salinity stress by maintaining sustained high rates of CEF and downregulating their short- term stress responses, through formation of a PSI-supercomplex.

ADP + P+

NADPH ATP H+ H+ H+ + + H+ H NADP H+ Stroma Fdx FNR

PQH2 e-

PQH2 LHCI P Cyt b6f LHCI

LHCII 680 LHCII P700 - - e e- e Lumen OEC PC H+ + H+ H+ Linear electron 3H 2H2O O2 + 4H+ flow

Photosystem II Cytochrome b6f Photosystem I ATP synthase ADP + P+

NADPH ATP H+ H+ H+ Cyclic electron + + H+ flow NADP H H+ Stroma Fdx FNR

PQH2

PQH2 LHCI P Cyt b6f LHCI

LHCII 680 LHCII P700 - - e e- e Lumen OEC PC + + H+ H H+ 3H 2H2O O2 + 4H+

Figure 1.1. Two modes of photosynthetic electron transport systems in photosynthetic organisms: Linear electron flow (Top) and Cyclic electron flow (Bottom). In LEF, electrons released from chlorophyll excitation and oxygen evolution at PSII, migrate towards PSI via the plastoquinone (PQ)

pool, cytb6f and plastocyanin (PC). A proton gradient is formed that leads to ATP production, while the electrons from PSI reduce NADP+ to NADPH through ferredoxin-NADP-reductase (FNR). On the other hand, under CEF promoting conditions, a supercomplex of PSI with other proteins can be formed. During

CEF, electrons cycle around PSI and cyt b6f via PQ pool and PC, generating proton pump that leads to ATP production. As electrons are cycling around PSI, no NADPH is produced.

Short-term Long-term SHORT-TERM ADAPTATION STRESS ACCLIMATION

TIME

sec min hour days weeks months years

Figure 1.2. Differences between short-term stress, acclimation and adaptation in green algae. Stress responses in photosynthetic organisms are dependent on the exposure time to the stress. Initially (seconds to minutes) the organism expresses short-term stress responses to decrease the excitation pressure on the photosynthetic apparatus (eg. state transitions). On the other hand, if the organism is exposed to stress for a longer period of time (hours/days or weeks/months), acclimation (short-term or long-term, respectively) strategies are employed (eg. changes in protein abundance, reorganization of photosynthetic apparatus). Last, if the organism is exposed to the stress for many years, adaptation occurs, where the organism permanently alters its genome to survive in the stress condition.

4.5 m ICE

Salinity dominant at 5 m LAKE WATER

dominant at 17 m ICE-MDV

700 mM NaCl at 17 m

40 m UWO241

0 PAR (µmol photons m-2 s-1) 50

Figure 1.3. A model of East Lake Bonney in MCM Dry Valleys. Depths at which the two psychrophilic Chlamydomonas sp. ICE-MDV and UWO241are abundant are shown. Salinity (black) increases with the depth whereas PAR (yellow) intensity decreases. PAR: Photosynthetic active radiation

1.6 REFERENCES

Aguilera, A., Souza-Egipsy, V., & Amils, R. (2012). Photosynthesis in Extreme Environments. Artificial Photosynthesis. https://doi.org/10.5772/26940

Airaki, M., Leterrier, M., Mateos, R. M., Valderrama, R., Chaki, M., Barroso, J. B., … Corpas, F. J. (2012). Metabolism of reactive oxygen species and reactive nitrogen species in pepper (Capsicum annuum L.) plants under low temperature stress. Plant, Cell and Environment, 35(2), 281–295. https://doi.org/10.1111/j.1365-3040.2011.02310.x

Alexandratos, N., & Brunismas, J. (2012). WORLD AGRICULTURE TOWARDS 2030/2050. JESA Working Papers 12-03. https://doi.org/10.22004/ag.econ.288998

Allakhverdiev, S. I., & Murata, N. (2008). Salt stress inhibits photosystems II and I in cyanobacteria. Photosynthesis Research, 98(1–3), 529–539. https://doi.org/10.1007/s11120- 008-9334-x

Alric, J. (2010). Cyclic electron flow around photosystem I in unicellular green algae. Photosynthesis Research, 106(1–2), 47–56. https://doi.org/10.1007/s11120-010-9566-4

Alric, J. (2015). The plastoquinone pool, poised for cyclic electron flow? Frontiers in Plant Science, 6(JULY), 1–4. https://doi.org/10.3389/fpls.2015.00540

Alric, J., Lavergne, J., & Rappaport, F. (2010). Redox and ATP control of photosynthetic cyclic electron flow in Chlamydomonas reinhardtii (I) aerobic conditions. Biochimica et Biophysica Acta - Bioenergetics, 1797(1), 44–51. https://doi.org/10.1016/j.bbabio.2009.07.009

Bai, X., Yang, L., Yang, Y., Ahmad, P., Yang, Y., & Hu, X. (2011). Deciphering the protective role of nitric oxide against salt stress at the physiological and proteomic levels in maize. Journal of Proteome Research, 10(10), 4349–4364. https://doi.org/10.1021/pr200333f

Bonente, G., Pippa, S., Castellano, S., Bassi, R., & Ballottari, M. (2012). Acclimation of Chlamydomonas reinhardtii to different growth irradiances. Journal of Biological Chemistry, 287(8), 5833–5847. https://doi.org/10.1074/jbc.M111.304279

Cardona, T., Shao, S., & Nixon, P. J. (2018). Enhancing photosynthesis in plants: the light reactions. Essays in Biochemistry, 62(1), 85–94. https://doi.org/10.1042/EBC20170015

Chang, R. L., Ghamsari, L., Manichaikul, A., Hom, E. F. Y., Balaji, S., Fu, W., … Papin, J. A. (2011). Metabolic network reconstruction of Chlamydomonas offers insight into light-driven algal metabolism. Molecular Systems Biology, 7(518). https://doi.org/10.1038/msb.2011.52

Chen, H., Hu, J., Qiao, Y., Chen, W., Rong, J., Zhang, Y., … Wang, Q. (2015). Ca 2+ -regulated cyclic electron flow supplies ATP for nitrogen starvation-induced lipid biosynthesis in green alga. Scientific Reports, 5(October), 1–12. https://doi.org/10.1038/srep15117

Chen, X., Tian, D., Kong, X., Chen, Q., Abd, A. A., Hu, X., & Jia, A. (2016). The role of nitric oxide signalling in response to salt stress in Chlamydomonas reinhardtii. Planta, 244(3), 651– 669. https://doi.org/10.1007/s00425-016-2528-0

Cook, G., Teufel, A., Kalra, I., Li, W., Wang, X., Priscu, J., & Morgan-Kiss, R. (2019). The Antarctic psychrophiles Chlamydomonas spp. UWO241 and ICE-MDV exhibit differential restructuring of photosystem I in response to iron. Photosynthesis Research, 141(2), 209– 228. https://doi.org/10.1007/s11120-019-00621-0

Cvetkovska, M., Smith, R., & Huner, N. P. A. (2017). Chilling out : the evolution and diversification of psychrophilic algae with a focus on Chlamydomonadales, 1169–1184. https://doi.org/10.1007/s00300-016-2045-4

Dietzel, L., Bräutigam, K., & Pfannschmidt, T. (2008). Photosynthetic acclimation: State transitions and adjustment of photosystem stoichiometry - Functional relationships between short-term and long-term light quality acclimation in plants. FEBS Journal, 275(6), 1080–

1088. https://doi.org/10.1111/j.1742-4658.2008.06264.x

Dijkhuizen, L., & Harder, W. (1985). Current views on the regulation of autotrophic carbon dioxide fixation via the Calvin cycle in bacteria. Antonie van Leeuwenhoek, 50(1984), 473– 487.

Dolhi, J. M., Maxwell, D. P., & Morgan-Kiss, R. M. (2013). Review: The Antarctic Chlamydomonas raudensis: An emerging model for cold adaptation of photosynthesis. Extremophiles, 17(5), 711–722. https://doi.org/10.1007/s00792-013-0571-3

Dolhi, J. M., Teufel, A. G., Kong, W., & Morgan-Kiss, R. M. (2015). Diversity and spatial distribution of autotrophic communities within and between ice-covered Antarctic lakes (McMurdo Dry Valleys). Limnology and Oceanography, 60(3), 977–991. https://doi.org/10.1002/lno.10071

Ebenhöh, O., Fucile, G., Finazzi, G., Rochaix, J. D., & Goldschmidt-Clermont, M. (2014). Short- term acclimation of the photosynthetic electron transfer chain to changing light: A mathematical model. Philosophical Transactions of the Royal Society B: Biological Sciences, 369(1640). https://doi.org/10.1098/rstb.2013.0223

Ensminger, I., Busch, F., & Huner, N. P. A. (2006). Photostasis and cold acclimation: Sensing low temperature through photosynthesis. Physiologia Plantarum, 126(1), 28–44. https://doi.org/10.1111/j.1399-3054.2006.00627.x

Erickson, E., Wakao, S., & Niyogi, K. K. (2015). Light stress and photoprotection in Chlamydomonas reinhardtii. Plant Journal, 82(3), 449–465. https://doi.org/10.1111/tpj.12825

Fang, W., Si, Y., Douglass, S., Casero, D., Merchant, S. S., Pellegrini, M., … Spaldinga, M. H. (2012). Transcriptome-wide changes in Chlamydomonas reinhardtii gene expression regulated by carbon dioxide and the CO2-concentrating mechanism regulator CIA5/CCM1. Plant Cell, 24(5), 1876–1893. https://doi.org/10.1105/tpc.112.097949

Finazzi, G., & Johnson, G. N. (2016). Cyclic electron flow: facts and hypotheses. Photosynthesis Research, 129(3), 227–230. https://doi.org/10.1007/s11120-016-0306-2

Fritsen, C. H., & Priscu, J. C. (1999). Seasonal change in the optical properties of the permanent ice cover on Lake Bonney, Antarctica: Consequences for lake productivity and phytoplankton dynamics. Limnology and Oceanography, 44(2), 447–454. https://doi.org/10.4319/lo.1999.44.2.0447

Girolomoni, L., Cazzaniga, S., Pinnola, A., Perozeni, F., Ballottari, M., & Bassi, R. (2019). LHCSR3 is a nonphotochemical quencher of both photosystems in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America, 116(10), 4212–4217. https://doi.org/10.1073/pnas.1809812116

Girolomoni, L., Ferrante, P., Berteotti, S., Giuliano, G., Bassi, R., & Ballottari, M. (2017). The function of LHCBM4/6/8 antenna proteins in Chlamydomonas reinhardtii. Journal of Experimental Botany, 68(3), 627–641. https://doi.org/10.1093/jxb/erw462

Govindjee, Shevela, D., & Björn, L. O. (2017). Evolution of the Z-scheme of photosynthesis: a perspective. Photosynthesis Research, 133(1–3), 5–15. https://doi.org/10.1007/s11120-016- 0333-z

Goyal, A. (2007a). Osmoregulation in Dunaliella, Part I: Effects of osmotic stress on photosynthesis, dark respiration and glycerol metabolism in Dunaliella tertiolecta and its salt- sensitive mutant (HL 25/8). Plant Physiology and Biochemistry, 45(9), 696–704. https://doi.org/10.1016/j.plaphy.2007.05.008

Goyal, A. (2007b). Osmoregulation in Dunaliella, Part II: Photosynthesis and starch contribute carbon for glycerol synthesis during a salt stress in Dunaliella tertiolecta. Plant Physiology and Biochemistry, 45(9), 705–710. https://doi.org/10.1016/j.plaphy.2007.05.009

Goyal, A., & Tolbert, N. E. (1990). Salicylhydroxamic acid (SHAM) inhibition of the dissolved inorganic carbon concentrating process in unicellular green algae. Plant Physiology, 92(3),

630–636. https://doi.org/10.1104/pp.92.3.630

Goyal, A., & Tolbert, N. E. (1996). Association of glycolate oxidation with photosynthetic electron transport in plant and algal chloroplasts. Proceedings of the National Academy of Sciences of the United States of America, 93(8), 3319–3324. https://doi.org/10.1073/pnas.93.8.3319

Gururani, M. A., Venkatesh, J., & Tran, L. S. P. (2015). Regulation of photosynthesis during abiotic stress-induced photoinhibition. Molecular Plant, 8(9), 1304–1320. https://doi.org/10.1016/j.molp.2015.05.005

He, Y., Fu, J., Yu, C., Wang, X., Jiang, Q., Hong, J., … Jiang, D. (2015). Increasing cyclic electron flow is related to Na+ sequestration into vacuoles for salt tolerance in soybean. Journal of Experimental Botany, 66(21), 6877–6889. https://doi.org/10.1093/jxb/erv392

Hema, R., Senthil-Kumar, M., Shivakumar, S., Chandrasekhara Reddy, P., & Udayakumar, M. (2007). Chlamydomonas reinhardtii, a model system for functional validation of abiotic stress responsive genes. Planta, 226(3), 655–670. https://doi.org/10.1007/s00425-007-0514-2

Ho, S. H., Nakanishi, A., Kato, Y., Yamasaki, H., Chang, J. S., Misawa, N., … Kondo, A. (2017). Dynamic metabolic profiling together with transcription analysis reveals salinity-induced starch-To-lipid biosynthesis in alga Chlamydomonas sp. JSC4. Scientific Reports, 7(October 2016), 1–7. https://doi.org/10.1038/srep45471

Huang, W., Fu, P. L., Jiang, Y. J., Zhang, J. L., Zhang, S. B., Hu, H., & Cao, K. F. (2013). Differences in the responses of photosystem I and photosystem II of three tree species Cleistanthus sumatranus, Celtis philippensis and Pistacia weinmannifolia exposed to a prolonged drought in a tropical limestone forest. Tree Physiology, 33(2), 211–220. https://doi.org/10.1093/treephys/tps132

Huang, W., Zhang, S. B., & Cao, K. F. (2010). Stimulation of cyclic electron flow during recovery after chilling-induced photoinhibition of PSII. Plant and Cell Physiology, 51(11), 1922–1928. https://doi.org/10.1093/pcp/pcq144

Huang, W., Zhang, S. B., Xu, J. C., & Liu, T. (2017). Plasticity in roles of cyclic electron flow around photosystem I at contrasting temperatures in the chilling-sensitive plant Calotropis

gigantea. Environmental and Experimental Botany, 141(May), 145–153. https://doi.org/10.1016/j.envexpbot.2017.07.011

Huang, Z., Liangliang, S., Wenda, W., Zhiyuan, M., Xiaohan, Y., Tingyun, K., … Guangye, H. (2021). Structure of photosystem I-LHCI-LHCII from the green alga Chlamydomonas reinhardtii in State 2. Nature Communications, (12(1)), 1–14. https://doi.org/10.1038/s41467- 021-21362-6

Hüner, N. P. A., Bode, R., Dahal, K., Hollis, L., Rosso, D., Krol, M., & Ivanov, A. G. (2012). Chloroplast redox imbalance governs phenotypic plasticity: The “grand design of photosynthesis” revisited. Frontiers in Plant Science, 3(NOV), 1–12. https://doi.org/10.3389/fpls.2012.00255

Huner, N. P. A., Öquist, G., & Sarhan, F. (1998). Energy balance and acclimation to light and cold, 1385(98).

Ivanov, A. G., Morgan, R. M., Gray, G. R., Velitchkova, M. Y., & Huner, N. P. A. (1998). Temperature/light dependent development of selective resistance to photoinhibition of photosystem I. FEBS Letters, 430(3), 288–292. https://doi.org/10.1016/S0014- 5793(98)00681-4

Iwai, M., Takizawa, K., Tokutsu, R., Okamuro, A., Takahashi, Y., & Minagawa, J. (2010). Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis. Nature, 464(7292), 1210–1213. https://doi.org/10.1038/nature08885

Jeffrey, S. W., & Humphrey, G. F. (1975). New spectrophotometric equations for determining chlorophylls a, b, c1 and c2 in higher plants, algae and natural phytoplankton. Biochemie Und Physiologie Der Pflanzen, 167(2), 191–194. https://doi.org/10.1016/s0015-3796(17)30778-3

Kalra, I., Wang, X., Cvetkovska, M., Jeong, J., McHargue, W., Zhang, R., … Morgan-Kissa, R. (2020). Chlamydomonas sp. UWO 241 exhibits high cyclic electron flow and rewired metabolism under high salinity. Plant Physiology, 183(2), 588–601. https://doi.org/10.1104/pp.19.01280

Kato, Y., Ho, S. H., Vavricka, C. J., Chang, J. S., Hasunuma, T., & Kondo, A. (2017). Evolutionary

engineering of salt-resistant Chlamydomonas sp. strains reveals salinity stress-activated starch-to-lipid biosynthesis switching. Bioresource Technology, 245, 1484–1490. https://doi.org/10.1016/j.biortech.2017.06.035

Khona, D. K., Shirolikar, S. M., Gawde, K. K., Hom, E., Deodhar, M. A., & D’Souza, J. S. (2016). Characterization of salt stress-induced palmelloids in the green alga, Chlamydomonas reinhardtii. Algal Research, 16(July), 434–448. https://doi.org/10.1016/j.algal.2016.03.035

Knopf, J. A., & Shapira, M. (2005). Degradation of Rubisco SSU during oxidative stress triggers aggregation of Rubisco particles in Chlamydomonas reinhardtii. Planta, 222(5), 787–793. https://doi.org/10.1007/s00425-005-0023-0

Kono, M., & Terashima, I. (2014). Long-term and short-term responses of the photosynthetic electron transport to fluctuating light. Journal of Photochemistry and Photobiology B: Biology, 137, 89–99. https://doi.org/10.1016/j.jphotobiol.2014.02.016

Kramer, D. M., & Evans, J. R. (2011). The importance of energy balance in improving photosynthetic productivity. Plant Physiology, 155(1), 70–78. https://doi.org/10.1104/pp.110.166652

Kumar, V., Wani, S. H., Penna, S., & Tran, L. S. P. (2018). Salinity Responses and Tolerance in Plants, Volume 1: Targeting Sensory, Transport and Signaling Mechanisms. Salinity Responses and Tolerance in Plants, Volume 1: Targeting Sensory, Transport and Signaling Mechanisms. https://doi.org/10.1007/978-3-319-75671-4

Kuo, E. Y. H., Chang, H. L., Lin, S. T., & Lee, T. M. (2020). High Light-Induced Nitric Oxide Production Induces Autophagy and Cell Death in Chlamydomonas reinhardtii. Frontiers in Plant Science, 11(June), 1–21. https://doi.org/10.3389/fpls.2020.00772

Laemmli, U. K. (1970). Cleavage of Structural Proteins during the Assembly of the Head of Bacteriophage T4. Nature, 227, 680–685.

Lemeille, S., & Rochaix, J. D. (2010). State transitions at the crossroad of thylakoid signalling pathways. Photosynthesis Research, 106(1–2), 33–46. https://doi.org/10.1007/s11120-010- 9538-8

León, R., & Galván, F. (1994). Halotolerance studies on Chlamydomonas reinhardtii: glycerol excretion by free and immobilized cells. Journal of Applied Phycology, 6(1), 13–20. https://doi.org/10.1007/BF02185898

Li, J., Tietz, S., Cruz, J. A., Strand, D. D., Xu, Y., Chen, J., … Hu, J. (2019). Photometric screens identified Arabidopsis peroxisome proteins that impact photosynthesis under dynamic light conditions. Plant Journal, 97(3), 460–474. https://doi.org/10.1111/tpj.14134

Li, W., & Morgan-Kiss, R. M. (2019). Influence of environmental drivers and potential interactions on the distribution of microbial communities from three permanently stratified Antarctic lakes. Frontiers in Microbiology, 10(MAY), 1–16. https://doi.org/10.3389/fmicb.2019.01067

Li, W., Podar, M., & Morgan-Kiss, R. M. (2016). Ultrastructural and single-cell-level characterization reveals metabolic versatility in a microbial eukaryote community from an ice-covered Antarctic lake. Applied and Environmental Microbiology, 82(12), 3659–3670. https://doi.org/10.1128/AEM.00478-16

Lima Neto, M. C., Cerqueira, J. V. A., da Cunha, J. R., Ribeiro, R. V., & Silveira, J. A. G. (2017). Cyclic electron flow, NPQ and photorespiration are crucial for the establishment of young plants of Ricinus communis and Jatropha curcas exposed to drought. Plant Biology, 19(4), 650–659. https://doi.org/10.1111/plb.12573

Liska, A. J., Shevchenko, A., Pick, U., & Katz, A. (2004). Enhanced photosynthesis and redox energy production contribute to salinity tolerance in Dunaliella as revealed by homology- based proteomics. Plant Physiology, 136(1), 2806–2817. https://doi.org/10.1104/pp.104.039438

Lizottel, M. P., & Prism, J. C. (1994). Natural fluorescence and quantum yields in vertically stationary phytoplankton from perennially ice‐covered lakes. Limnology and Oceanography, 39(6), 1399–1410. https://doi.org/10.4319/lo.1994.39.6.1399

Lucker, B., & Kramer, D. M. (2013). Regulation of cyclic electron flow in Chlamydomonas reinhardtii under fluctuating carbon availability. Photosynthesis Research, 117(1–3), 449–

459. https://doi.org/10.1007/s11120-013-9932-0

Magneschi, L., Catalanotti, C., Subramanian, V., Dubini, A., Yang, W., Mus, F., … Grossman, A. R. (2012). A mutant in the ADH1 gene of Chlamydomonas reinhardtii elicits metabolic restructuring during anaerobiosis. Plant Physiology, 158(3), 1293–1305. https://doi.org/10.1104/pp.111.191569

Melis, A. (1991). Dynamics of photosynthetic membrane composition and function. BBA - Bioenergetics, 1058(2), 87–106. https://doi.org/10.1016/S0005-2728(05)80225-7

Minagawa, J. (2016). A Supercomplex of Cytochrome bf and Photosystem I for Cyclic Electron Flow, 453–462. https://doi.org/10.1007/978-94-017-7481-9_23

Morgan-Kiss, R., Ivanov, A. G., Williams, J., Khan, M., & Huner, N. P. A. (2002). Differential thermal effects on the energy distribution between photosystem II and photosystem I in thylakoid membranes of a psychrophilic and a mesophilic alga. Biochimica et Biophysica Acta - Biomembranes, 1561(2), 251–265. https://doi.org/10.1016/S0005-2736(02)00352-8

Morgan-Kiss, R. M., Ivanov, A. G., & Huner, N. P. A. (2002). The Antarctic psychrophile, Chlamydomonas subcaudata, is deficient in State I-State II transitions. Planta, 214(3), 435– 445. https://doi.org/10.1007/s004250100635

Morgan-Kiss, R. M., Ivanov, A. G., Modla, S., Czymmek, K., Hüner, N. P. A., Priscu, J. C., … Hanson, T. E. (2008). Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp., strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles, 12(5), 701–711. https://doi.org/10.1007/s00792-008-0176-4

Morgan-Kiss, R. M., Priscu, J. C., Pocock, T., Gudynaite-Savitch, L., & Huner, N. P. A. (2006). Adaptation and Acclimation of Photosynthetic Microorganisms to Permanently Cold Environments. Microbiology and Molecular Biology Reviews, 70(1), 222–252. https://doi.org/10.1128/mmbr.70.1.222-252.2006

Morgan, R. M., Ivanov, A. G., Priscu, J. C., Maxwell, D. P., & Huner, N. P. A. (1998). Structure and composition of the photochemical apparatus of the Antarctic green alga, Chlamydomonas subcaudata. Photosynthesis Research, 56(3), 303–314.

https://doi.org/10.1023/A:1006048519302

Morton, M. J. L., Awlia, M., Al-Tamimi, N., Saade, S., Pailles, Y., Negrão, S., & Tester, M. (2019). Salt stress under the scalpel – dissecting the genetics of salt tolerance. Plant Journal, 97(1), 148–163. https://doi.org/10.1111/tpj.14189

Mou, S., Zhang, X., Ye, N., Dong, M., Liang, C., Liang, Q., … Zheng, Z. (2012). Cloning and expression analysis of two different LhcSR genes involved in stress adaptation in an Antarctic microalga, Chlamydomonas sp. ICE-L. Extremophiles, 16(2), 193–203. https://doi.org/10.1007/s00792-011-0419-7

Müller, P., Li, X. P., & Niyogi, K. K. (2001). Non-photochemical quenching. A response to excess light energy. Plant Physiology, 125(4), 1558–1566. https://doi.org/10.1104/pp.125.4.1558

Munekage, Y., Hojo, M., Meurer, J., Endo, T., Tasaka, M., & Shikanai, T. (2002). PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in Arabidopsis. Cell, 110(3), 361–371. https://doi.org/10.1016/S0092-8674(02)00867-X

Nawrocki, W. J., Liu, X., & Croce, R. (2020). Chlamydomonas reinhardtii exhibits de facto constitutive NPQ capacity in physiologically relevant conditions. Plant Physiology, 182(1), 472–479. https://doi.org/10.1104/pp.19.00658

Neale, P. J., & Melis, A. (1989). Salinity-stress Enhances Photoinhibition of Photosynthesis in Chlamydomonas reinhardtii. Journal of Plant Physiology, 134(5), 619–622. https://doi.org/10.1016/S0176-1617(89)80158-0

Neelam, S., & Subramanyam, R. (2013). Alteration of photochemistry and protein degradation of photosystem II from Chlamydomonas reinhardtii under high salt grown cells. Journal of Photochemistry and Photobiology B: Biology, 124, 63–70. https://doi.org/10.1016/j.jphotobiol.2013.04.007

Negrão, S., Schmöckel, S. M., & Tester, M. (2017). Evaluating physiological responses of plants to salinity stress. Annals of Botany, 119(1), 1–11. https://doi.org/10.1093/aob/mcw191

Oren, A. (2014). The ecology of Dunaliella in high-salt environments. Journal of Biological

Research (Greece), 21(1), 1–8. https://doi.org/10.1186/s40709-014-0023-y

Perrineau, M. M., Zelzion, E., Gross, J., Price, D. C., Boyd, J., & Bhattacharya, D. (2014). Evolution of salt tolerance in a laboratory reared population of Chlamydomonas reinhardtii. Environmental Microbiology, 16(6), 1755–1766. https://doi.org/10.1111/1462-2920.12372

Petroutsos, D., Busch, A., Janßen, I., Trompelt, K., Bergner, S. V., Weinl, S., … Hippler, M. (2011). The chloroplast calcium sensor CAS is required for photoacclimation in Chlamydomonas reinhardtii. Plant Cell, 23(8), 2950–2963. https://doi.org/10.1105/tpc.111.087973

Petroutsos, D., Tokutsu, R., Maruyama, S., Flori, S., Greiner, A., Magneschi, L., … Minagawa, J. (2016). A blue-light photoreceptor mediates the feedback regulation of photosynthesis. Nature, 537(7621), 563–566. https://doi.org/10.1038/nature19358

Plouviez, M., Wheeler, D., Shilton, A., Packer, M. A., McLenachan, P. A., Sanz-Luque, E., … Guieysse, B. (2017). The biosynthesis of nitrous oxide in the green alga Chlamydomonas reinhardtii. Plant Journal, 91(1), 45–56. https://doi.org/10.1111/tpj.13544

Priscu, J. C. (1998). Ecosystem Dynamics in a Polar Desert, 72.

Priscu, J. C., & Spigel, R. H. (1996). Evolution of temperature and salt structure of Lake Bonney, a chemically stratified Antarctic lake. Hydrobiologia, 321(3), 177–190.

Priscu, J. C., Wolf, F., Takacs, D., & Fritsen, H. (1999). Carbon Transformatiin a Perennially Ice- Covered Antarctic Lake, 49(12), 997–1008.

Raymond, J. A., & Morgan-Kiss, R. (2013). Separate Origins of Ice-Binding Proteins in Antarctic Chlamydomonas Species. PLoS ONE, 8(3), 6–11. https://doi.org/10.1371/journal.pone.0059186

Raymond, J. A., Morgan-Kiss, R., & Stahl-Rommel, S. (2020). Glycerol Is an Osmoprotectant in Two Antarctic Chlamydomonas Species From an Ice-Covered Saline Lake and Is Synthesized by an Unusual Bidomain Enzyme. Frontiers in Plant Science, 11(August), 1–8. https://doi.org/10.3389/fpls.2020.01259

Reynoso, G. T., & de Gamboa, B. A. (1982). Salt tolerance in the freshwater algae Chlamydomon as reinhardii: Effect of proline and taurine. Comparative Biochemistry and Physiology -- Part A: Physiology, 73(1), 95–99. https://doi.org/10.1016/0300-9629(82)90098-6

Roach, T., & Krieger-Liszkay, A. (2014). Regulation of Photosynthetic Electron Transport and Photoinhibition. Current Protein & Peptide Science, 15(4), 351–362. https://doi.org/10.2174/1389203715666140327105143

Rochaix, J. D. (2007). Role of thylakoid protein kinases in photosynthetic acclimation. FEBS Letters, 581(15), 2768–2775. https://doi.org/10.1016/j.febslet.2007.04.038

Sabeeha S. Merchant1,*, Simon E. Prochnik2,*, Olivier Vallon3, Elizabeth H. Harris4, Steven A. Sanderfoot11, Martin H. Spalding12, Vladimir V. Kapitonov13, Qinghu Ren14, Patrick Laurence Maréchal-Drouard7, Wallace F. Marshall8, Liang-Hu Qu9, David R. Nels, A. (2007). The Chlamydomonas Genome Reveals the Evolution of Key. National Institutes of Health, 318(5848), 245–250.

Saroussi, S. I., Wittkopp, T. M., & Grossman, A. R. (2016). The Type II NADPH dehydrogenase facilitates cyclic electron flow, energy-dependent quenching, and chlororespiratory metabolism during acclimation of Chlamydomonas reinhardtii to nitrogen deprivation. Plant Physiology, 170(4), 1975–1988. https://doi.org/10.1104/pp.15.02014

Shetty, P., Gitau, M. M., & Maróti, G. (2019). Salinity Stress Responses and Adaptation Mechanisms in Eukaryotic Green Microalgae. Cells, 8(12), 1–16. https://doi.org/10.3390/cells8121657

Shoun, H., Fushinobu, S., Jiang, L., Kim, S. W., & Wakagi, T. (2012). Fungal denitrification and nitric oxide reductase cytochrome P450nor. Philosophical Transactions of the Royal Society B: Biological Sciences, 367(1593), 1186–1194. https://doi.org/10.1098/rstb.2011.0335

Sithtisarn, S., Yokthongwattana, K., Mahong, B., Roytrakul, S., Paemanee, A., Phaonakrop, N., & Yokthongwattana, C. (2017). Comparative proteomic analysis of Chlamydomonas reinhardtii control and a salinity-tolerant strain revealed a differential protein expression pattern. Planta, 246(5), 843–856. https://doi.org/10.1007/s00425-017-2734-4

Sivakumar, P., Sharmila, P., & Pardha Saradhi, P. (2000). Proline alleviates salt-stress-induced enhancement in ribulose-1,5-bisphosphate oxygenase activity. Biochemical and Biophysical Research Communications, 279(2), 512–515. https://doi.org/10.1006/bbrc.2000.4005

Steinbeck, J., Ross, I. L., Rothnagel, R., Gäbelein, P., Schulze, S., Giles, N., … Hankamer, B. (2018). Structure of a PSI-LHCI-cyt b6f supercomplex in Chlamydomonas reinhardtii promoting cyclic electron flow under anaerobic conditions. Proceedings of the National Academy of Sciences of the United States of America, 115(41), 10517–10522. https://doi.org/10.1073/pnas.1809973115

Strand, D. D., Livingston, A. K., Satoh-Cruz, M., Froehlich, J. E., Maurino, V. G., & Kramer, D. M. (2015). Activation of cyclic electron flow by hydrogen peroxide in vivo. Proceedings of the National Academy of Sciences of the United States of America, 112(17), 5539–5544. https://doi.org/10.1073/pnas.1418223112

Subramanyam, R., Jolley, C., Thangaraj, B., Nellaepalli, S., Webber, A. N., & Fromme, P. (2010). Structural and functional changes of PSI-LHCI supercomplexes of Chlamydomonas reinhardtii cells grown under high salt conditions. Planta, 231(4), 913–922. https://doi.org/10.1007/s00425-009-1097-x

Sudhir, P., & Murthy, S. D. S. (2004). Effects of salt stress on basic processes of photosynthesis. Photosynthetica, 42(4), 481–486. https://doi.org/10.1007/S11099-005-0001-6

Suorsa, M. (2015). Cyclic electron flow provides acclimatory plasticity for the photosynthetic machinery under various environmental conditions and developmental stages. Frontiers in Plant Science, 6(September), 1–8. https://doi.org/10.3389/fpls.2015.00800

Szyszka-Mroz, B., Cvetkovska, M., Ivanov, A. G., Smith, D. R., Possmayer, M., Maxwell, D. P., & Hüner, N. P. A. (2019). Cold-adapted protein kinases and thylakoid remodeling impact energy distribution in an antarctic psychrophile. Plant Physiology, 180(3), 1291–1309. https://doi.org/10.1104/pp.19.00411

Szyszka-Mroz, B., Pittock, P., Ivanov, A. G., Lajoie, G., & Hüner, N. P. A. (2015). The antarctic psychrophile Chlamydomonas sp. UWO 241 preferentially phosphorylates a photosystem I-

cytochrome b6/f supercomplex. Plant Physiology, 169(1), 717–736. https://doi.org/10.1104/pp.15.00625

Szyszka, B., Ivanov, A. G., & Hüner, N. P. A. (2007). Psychrophily is associated with differential energy partitioning, photosystem stoichiometry and polypeptide phosphorylation in Chlamydomonas raudensis. Biochimica et Biophysica Acta - Bioenergetics, 1767(6), 789– 800. https://doi.org/10.1016/j.bbabio.2006.12.001

Takahashi, H., Clowez, S., Wollman, F. A., Vallon, O., & Rappaport, F. (2013). Cyclic electron flow is redox-controlled but independent of state transition. Nature Communications, 4. https://doi.org/10.1038/ncomms2954

Takahashi, H., Iwai, M., Takahashi, Y., & Minagawa, J. (2006). Identification of the mobile light- harvesting complex II polypeptides for state transitions in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America, 103(2), 477–482. https://doi.org/10.1073/pnas.0509952103

Terashima, M., Petroutsos, D., Hüdig, M., Tolstygina, I., Trompelt, K., Gäbelein, P., … Hippler, M. (2012). Calcium-dependent regulation of cyclic photosynthetic electron transfer by a CAS, ANR1, and PGRL1 complex. Proceedings of the National Academy of Sciences of the United States of America, 109(43), 17717–17722. https://doi.org/10.1073/pnas.1207118109

Valledor, Luis; Weckwerth, W. (2014). Chapter 25. In Plant Proteomics: Methods and Protocols, Methods in Molecular Biology, (Vol. 1072, pp. 347–358). https://doi.org/10.1016/b978-0-08- 011049-3.50031-5

Van Lis, R., Popek, M., Couté, Y., Kosta, A., Drapier, D., Nitschke, W., & Atteia, A. (2017). Concerted up-regulation of aldehyde/alcohol dehydrogenase (ADHE) and starch in Chlamydomonas reinhardtii increases survival under dark anoxia. Journal of Biological Chemistry, 292(6), 2395–2410. https://doi.org/10.1074/jbc.M116.766048

Voss, I., Sunil, B., Scheibe, R., & Raghavendra, A. S. (2013). Emerging concept for the role of photorespiration as an important part of abiotic stress response. Plant Biology, 15(4), 713– 722. https://doi.org/10.1111/j.1438-8677.2012.00710.x

Wang, N., Qian, Z., Luo, M., Fan, S., Zhang, X., & Zhang, L. (2018). Identification of salt stress responding genes using transcriptome analysis in green alga Chlamydomonas reinhardtii. International Journal of Molecular Sciences, 19(11). https://doi.org/10.3390/ijms19113359

Wang, X., Liu, W., Xin, C., Zheng, Y., Cheng, Y., Sun, S., … Yuan, J. S. (2016). Enhanced limonene production in cyanobacteria reveals photosynthesis limitations. Proceedings of the National Academy of Sciences of the United States of America, 113(50), 14225–14230. https://doi.org/10.1073/pnas.1613340113

Wang, Y., Stessman, D. J., & Spalding, M. H. (2015). The CO2 concentrating mechanism and photosynthetic carbon assimilation in limiting CO2: How Chlamydomonas works against the gradient. Plant Journal, 82(3), 429–448. https://doi.org/10.1111/tpj.12829

Wingler, A., Lea, P. J., Quick, W. P., & Leegood, R. C. (2000). Photorespiration: Metabolic pathways and their role in stress protection. Philosophical Transactions of the Royal Society B: Biological Sciences, 355(1402), 1517–1529. https://doi.org/10.1098/rstb.2000.0712

Yang, X., Li, Y., Chen, H., Huang, J., Zhang, Y., Qi, M., … Li, T. (2020). Photosynthetic response mechanism of soil salinity-induced cross-tolerance to subsequent drought stress in tomato plants. Plants, 9(3), 1–15. https://doi.org/10.3390/plants9030363

Yi, S. Y., Ku, S. S., Sim, H. J., Kim, S. K., Park, J. H., Lyu, J. Il, … Liu, J. R. (2017). An alcohol dehydrogenase gene from synechocystis sp. Confers salt tolerance in transgenic tobacco. Frontiers in Plant Science, 8(November), 1–13. https://doi.org/10.3389/fpls.2017.01965

Ziotti, A. B. S., Silva, B. P., Sershen, & Lima Neto, M. C. (2019). Photorespiration is crucial for salinity acclimation in castor bean. Environmental and Experimental Botany, 167(May),

103845. https://doi.org/10.1016/j.envexpbot.2019.103845

CHAPTER 2

Chlamydomonas sp. UWO241 exhibits constitutively high cyclic electron flow and rewired metabolism under high salinity.

Isha Kalra, Xin Wang, Marina Cvetkovska, Jooyeon Jeong, William McHargue, Ru Zhang, Norman Hüner, Joshua S. Yuan, and Rachael Morgan-Kiss

Most of this chapter appeared in Kalra, I., Wang, X., Cvetkovska, M., Jeong, J., McHargue, W., Zhang, R., ... & Morgan-Kiss, R. (2020). Chlamydomonas sp. UWO 241 exhibits high cyclic electron flow and rewired metabolism under high salinity. Plant physiology, 183(2), 588-601.

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CHAPTER II. Chlamydomonas sp. UWO241 exhibits constitutively high cyclic

electron flow and rewired metabolism under high salinity.

2.1 INTRODUCTION

During photosynthesis light is transduced into stored energy through two major pathways, linear electron flow (LEF) and cyclic electron flow (CEF). In addition to satisfying the ATP shortage for efficient carbon fixation (Kramer and Evans, 2011), CEF-generated ATP may be used for

other energy-requiring processes, such as the CO2-concentrating mechanism of C4 photosynthesis (Takabayashi et al., 2005; Ishikawa et al., 2016), N2 fixation in cyanobacteria heterocysts (Magnuson et al., 2011; Magnuson and Cardona, 2016), and survival under environmental stress (Suorsa, 2015). In addition, CEF is utilized under short-term stress by rapidly inducing a transthylakoid pH change and triggering nonphotochemical energy dissipation (Lucker and Kramer, 2013; Yamori et al., 2016b). Previously, it was assumed that CEF was dependent upon state transitions (Iwai et al., 2010); however, recent studies suggest that CEF induction is independent of state transitions and is sensitive to other signals (Takahashi et al., 2013; Strand et al., 2015). Formation of protein supercomplexes has been associated with CEF initiation (Minagawa, 2016). The first supercomplex was isolated by Iwai and colleagues (Iwai et al., 2010) in Chlamydomonas reinhardtii under dark/anaerobiosis. The supercomplex is composed of

PSI, LHCII, cytochrome b6f, PGRL1 and FNR (Ferredoxin NADP Reductase) (Iwai et al., 2010). Takahashi and colleagues (2013) identified another supercomplex in C. reinhardtii that is formed under conditions of anoxia and is regulated through the calcium sensing protein, CAS. Recently, the structure of the C. reinhardtii PSI supercomplex was solved which showed that dissociation of specific LHCI proteins (Lhca2 and Lhca9) are necessary prior to PSI supercomplex formation (Steinbeck et al., 2018). Around the globe, there are communities of photosynthetic organisms that have adapted to capture light energy and fix carbon under environmental conditions which are untenable for most model plant and algal species, including permanent low temperature environments in the Arctic and Antarctic (Dolhi et al., 2013; Morgan-Kiss et al., 2006). The McMurdo (MCM) Dry Valleys form the largest polar desert on the Antarctic continent and harbor numerous

permanently ice-covered lakes (Priscu et al., 1999; Morgan-Kiss et al., 2006). Microalgal communities are adapted to extreme conditions, including low temperatures, nutrient deficiency, super-saturated oxygen levels, and hypersalinity (Morgan-Kiss et al., 2006). Chlamydomonas sp. UWO241 (UWO241) was isolated from Lake Bonney in the 1990s (Neale and Priscu, 1990; 1995). In its native environment, UWO241 is exposed to year-round low temperatures (0°– 5°C), hypersalinity (700 mM NaCl), and extreme shade (<20 μmol photons m-2 s-1) of a narrow spectral range (350 – 450 nm). In early studies, it was reported that UWO241 appeared to exhibit permanent downregulation of PSI, estimated by a weak P700 photooxidation and an absence of a discernable PSI Chl a low temperature (77K) fluorescence emission peak under a range of treatments (Morgan-Kiss et al., 2002a; 2005; Szyszka et al., 2007; Cook et al., 2019). Morgan-Kiss et al. (2002b) reported that UWO241 lacks phosphorylation of LHCII and state transitions. Recently, Szyszka-Mroz et al. (2019) showed that UWO241 exhibits some LHCII phosphorylation and cold-adapted forms of the thylakoid protein kinases, STT7 and Stl1. They suggested that UWO241 may rely on energy spillover between PSII and PSI rather than state transitions (Szyska-Mroz et al., 2019). UWO241 maintains sustained CEF under steady-state growth conditions which is associated with a PSI supercomplex (Szyszka-Mroz et al., 2015; Cook et al., 2019). Formation of the UWO241 PSI supercomplex was present only in cultures acclimated to high salinity (700 mM NaCl) and its stability was disrupted in the presence of the kinase inhibitor, stauroporine (Szyszka-Mroz et al., 2015). These previous studies suggest a role for sustained CEF activity in UWO241 to survive long-term exposure to high salinity; however, the specific benefits of which are not understood. Here we investigated whether the outcomes of CEF provide UWO241 with constitutive photoprotection ability or energy generation. Through electrochomic shift and fluorescence spectroscopy, we identified increased capacity for qE and higher proton flux through ATP synthase in UWO241 under long-term salinity condition. Next, we conducted comparative whole cell proteomics and metabolomics to understand the effect of these CEF associated changes on downstream metabolism of UWO241. Our results indicate CEF under long-term acclimation to high salinity is associated with rewiring of primary carbon metabolism.

2.2 MATERIALS AND METHODS

2.2.1 Culture conditions, growth physiology.

Chlamydomonas sp. UWO241 (UWO241; CCMP1619) was grown in either Bold’s Basal Media (BBM, 0.43 mM NaCl) (Low salt, LS) or BBM supplemented with 700 mM NaCl (High salt, HS). Based on earlier studies (Morgan et al. 1998), UWO241 cultures were grown under a temperature/irradiance regime of 8oC/50 photons μmol m-2s-1. C.reinhardtii UTEX 90 was grown in BBM (LS) at 20°C/100 μmol photons m-2s-1. All cultures were grown in 250 ml glass pyrex tubes in temperature regulated aquaria under a 24-hour light cycle and were continuously aerated with sterile air supplied by aquarium pumps (Morgan-Kiss et al., 2008). Growth was monitored daily by optical density at wavelength of 750 nm. Maximum growth rates were calculated using natural log transformation of the optical density values during the exponential phase. Three biological replicates were performed, and all subsequent experiments were conducted on log-phase cultures. Oxygen evolution was measured at 8°C with Chlorolab 2 (Hansatech, UK) based on a Clark-type oxygen electrode, following the method described in Jeong et al. (2017). Cellular ATP and starch were determined using commercial kits, an ATP colometric/fluorometric assay kit (Catalogue No. K354, BioVision Inc.) and a starch assay kit (Catalogue No. SA20, Sigma-Aldrich), respectively. For supercomplex isolation, C. reinhardtii cells were exposed to either state 1 or state 2 conditions. For state 1 induction, mid-log phase cells were incubated in dark for 10 min to completely oxidize the PQ pool. For state 2 induction, mid-log phase cells were incubated in the dark under anaerobic conditions (N2 bubbling) for 30 min.

2.2.2 Low temperature (77K) fluorescence spectra.

Low temperature Chl a fluorescence emission spectra of whole cells and isolated Chl-protein complexes were measured using Luminescence Spectrometer LS50B (Perkin Elmer, USA) as described in Morgan et al. (2008) at 436 nm and 5 (isolate complexes) or 8 nm (whole cells) slit widths. Prior to the measurement, cultures were dark adapted for 10 min. Decompositional analysis was performed using a non-linear least squares algorithm using Microcal OriginPro Version 8.5.1 (Microcal Origin Northampton, MA). The fitting parameters for the Gaussian components (position, area and full width half-maximum, FWHM) were free running parameters.

2.2.3 P700 oxidation-reduction and cyclic electron flow.

Far red light induced photooxidation of P700 was used to determine rates of CEF as described by Morgan-Kiss et al. (2002b). A volume of exponential phase cultures representing 25 μg Chl a was dark adapted for 10 min and then filtered onto 25 mm GF/C filters (Whatman). Filters were measured on the Dual-PAM 100 instrument using the leaf attachment. The proportion of photooxidizable P700 was determined by monitoring absorbance changes at 820 nm and

expressed as the parameter (∆A820/A820). The signal was balanced, and the measuring light switched on. Far red (FR) light (λmax=715 nm, 10 Wm−2, Scott filter RG 715) was then switched on to oxidize P700. After steady-state oxidation levels were reached, the FR light was + red switched off to re-reduce P700. The half time for the reduction of P700 to P700 (t½ ) was calculated after the FR light was turned off as an estimate of relative rates of PSI-driven CEF (Ivanov et al., 1998). The re-reduction time for P700 was calculated using MicrocalTM OriginTM software (Microcal Software Inc., Northampton, MA, USA).

2.2.4 In vivo spectroscopy measurements.

Saturation-pulse chlorophyll fluorescence yield changes and dark interval relaxation kinetics (DIRK) of ECS were measured at 8°C with the IDEA spectrophotometer as described previously with some modifications (Sacksteder and Kramer, 2000; Zhang et al., 2009). A 2.5 mL of cell supplemented with 25 μL of 0.5 M NaHCO3 was pre-incubated in the dark for 10 min and followed by 10 min illumination of far-red light. The chlorophyll fluorescence and ECS were measured with the cells acclimated for 5 min in various actinic light intensities provided by red

LEDs. The PSII operating efficiency (ΦPSII) was calculated as Fq’/Fm’, NPQ as (Fm-Fm’)/Fm’. The

linear electron transport (LEF) was calculated from following equation: LEF = A × (fractionPSII)

× I × ΦPSII, where I is the light intensity, A is the absorptivity of the sample, which is generally

assumed to be 0.84 and fractionPSII is the fraction of absorbed light stimulating PSII (Baker,

2008). The fractionPSII of UWO241 grown in low-salt and high salt, measured by 77K fluorescence spectra, were 0.709 and 0.746, respectively. The total amplitude of the ECS signal

(ECSt) was used to estimate the proton motive force (pmf). The aggregate conductivity of the

thylakoid membrane to protons (gH+) was estimated from the inverse of lifetime of the rapid decay of ECS (τECS) (Baker et al., 2007). All ECS signals were normalized to the rapid rise in

ECS induced by a single turnover flash to account for changes in pigmentation (Livingston et al. 2010).

2.2.6 Thylakoid isolation.

Thylakoids were isolated according to Morgan-Kiss et al. (1998). Mid-log phase cultures were collected by centrifugation at 2500g for 5 min at 4°C. All buffers were kept ice-cold and contained 1 mM Pefabloc Sc (Sigma, USA) and 20 mM NaF. The pellet was resuspended in grinding buffer (0.3 M sorbitol, 10 mM NaCl, 5 mM MgCl2, 5 mM MgCl2, 1 mM benzamidine, 1mM amino-caproic acid). The cells were disrupted using chilled French press at 10,000 lb/in2 twice, and then and centrifuged at 23,700g for 30 min. The thylakoid pellet was resuspended in wash buffer (50 mM Tricine-NaOH [pH 7.8], 10 mM NaCl, 5 mM MgCl2) and centrifuged at 13,300xg for 20 min. The pellet was resuspended in storage buffer (0.3 M sorbitol, 10% glycerol, 50 mM Tricine-NaOH [pH 7.8], 10 mM NaCl and 5 mM MgCl2) and stored at -80°C until analysis.

2.2.7 SDS-PAGE and Immunoblotting.

SDS-PAGE was performed using Bio-Rad Mini-Protean system and 12% Urea-SDS gel (Laemmli, 1970). Thylakoid membranes were denatured using 50 mM DTT and incubated at 70°C for 5 min. Samples were loaded on equal protein basis (10 μg total protein). Proteins were transferred to nitrocellulose membrane using cold-wet transfer at 100 V for 2.5 hours. The membrane was blocked with TBST (Tris Buffer Saline Tween) buffer with 5% milk (Carnation). A primary antibody against PsaA (Cat No. AS06-172; Agrisera, Sweden) was used at 1:1000 dilution to probe for major reaction center protein of PSI. Membranes were then exposed to Protein A conjugated to horseradish peroxidase and blots were detected with ECL SelectTM Western Blotting Detection Reagent (Amersham).

2.2.8 Supercomplex isolation.

Sucrose step density centrifugation was used to isolate supercomplexes from exponentially grown cultures according to Szyszka-Mroz et al. (2015) with some modifications. Every step was performed in darkness and on ice. All buffers contained phosphatase (20 mM NaF) and protease (1 mM Pefabloc SC) inhibitor. Cells were collected by centrifugation and the pellet was washed twice in Buffer 1 (0.3 M Sucrose, 25 mM Hepes-KOH [pH 7.5], 1mM MgCl2). Cells

38 were disrupted using French press, as described above and broken cells were spun down at 50,000g for 30 min. The pellet was resuspended in Buffer 2 (0.3 M Sucrose, 5 mM Hepes-KOH [pH 7.5], 10 mM EDTA) and centrifuged at 50,000g for 30 min. The thylakoid pellet was resuspended gently in Buffer 3 (1.8 M Sucrose, 5mM Hepes-KOH [pH 7.5], 10 mM EDTA) and transferred to Ultra-clear tube (Catalogue No., 344060, Beckman Coulter, USA). The thylakoid prep was overlayed with Buffer 4 (1.3 M Sucrose, 5mM Hepes-KOH [pH 7.5], 10 mM EDTA) followed by Buffer 5 (0.5 M Sucrose, 5mM Hepes-KOH [pH 7.5]). This sucrose step gradient was ultra-centrifuged at 288,000g for 1 hour at 4°C using Sw40Ti rotor (Beckman coulter, USA). Purified thylakoids were collected and diluted (3-fold) in Buffer 6 (5 mM Hepes-KOH [pH 7.5], 10 mM EDTA) and centrifuged at 50,000xg to pellet the membrane. Linear sucrose gradients were made using freeze thaw method with Buffer 7a (1.3 M Sucrose, 5 mM Hepes- KOH [pH 7.5], 0.05% α-DDM) and Buffer 7b (0.1 M Sucrose, 5 mM Hepes-KOH [pH 7.5], 0.05% α-DDM). Briefly, two dilutions of Buffers 7a and 7b were made, Buffer 7-1 (2x Buffer 7a + 1x Buffer 7b) and Buffer 7-2 (1x Buffer 7 a + 2x Buffer 7b). To make the gradient, first 3 ml of Buffer 7a was poured into 12 ml ultra-clear tubes followed by flash freezing in liquid nitrogen. Next, Buffer 7-1 was poured on top, followed by flash freezing. This was repeated for Buffer 7-2 and Buffer 7b respectively. The frozen gradients were kept at 4°C overnight to thaw. For supercomplex isolation, thylakoid membranes (0.4 mg Chl) were resuspended in 1% n- dodecyl-alpha-maltoside (α-DDM) (Catalogue number D99020, Glycon Biochemicals, Germany) and incubated on ice in the dark for 25 min. Membranes were spun down to remove insoluble material and loaded onto a linear sucrose gradient described above (0.1 – 1.3 M sucrose) containing 0.05% α-DDM. Gradients were centrifuged at 288,000g for 21 hours at 4°C using SW40Ti rotor (Beckman Coulter, USA). Protein complexes were extracted using a 21- gauge needle.

2.2.9 Sample preparation for proteomics.

Whole cell proteins were extracted as described previously (Valledor and Weckwerth, 2014). Mid-log phase cells were collected by centrifugation at 2500g for 5 min (50 mg wet weight). The cell pellets were resuspended in an extraction buffer containing 100 mM Tris-HCl (pH 8.0), 10% (v/v) glycerol, 2 mM Pefabloc Sc, 10 mM DTT, and 1.2 % (v/v) plant protease inhibitor cocktail (Cat. No. P9599, Sigma). Samples were transferred to 2 mL screw cap tubes containing 25 mg of

39 zirconia beads (Cat. No. A6758, Biorad) and homogenized 3 times for 45 seconds in a BeadBeater (BioSpec). 20% SDS solution was added to the tubes and samples were incubated for 5 min at 95°C. The denatured proteins were centrifuged at 12,000g to pellet any insoluble material. Protein pellets were resuspended in 1.5 ml Tris buffer (50mM Tris-HCl, pH 8.0) containing 0.02% n-dodecyl-beta-maltoside (Glycon Biochemicals, Germany) and supplemented with 1X Halt™ protease and phosphatase inhibitor cocktail (Thermo-Scientific, Rockford, IL). After the protein extraction, the sample preparation for proteomics were conducted following our previously published method (Wang et al., 2016). Specifically, 100 μg of total protein were treated with 8 M Urea/5 mM DTT for 1 hour at 37°C, followed by alkylation with 15 mM iodoacetamide in dark for 30 minutes at room temperature. Samples were then diluted 4-folds with 50 mM Tris-HCl buffer and digested using Mass-spectrometry Grade Trypsin Gold (Promega, Madison, WI) at 1:100 w/w concentration for 16.5 hours at 37°C. The digested samples were cleaned using Sep-Pak C18 plus desalting columns (Waters Corporation, Milford, MA).

2.2.10 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS)

The whole cell proteomics were conducted using the Multidimensional Protein Identification Technology (MudPIT) based shotgun proteomics by loading digested peptides onto a biphasic strong cation exchange/reversed phase capillary column. The two dimensional (2D)-LC-MS/MS was conducted on an LTQ ion trap mass spectrometer (Thermo Finnegan, San Jose, CA) operated in the data-dependent acquisition mode. The full mass spectra were recorded at 300- 1700 m/z, and the 5 most abundant peaks of each scan were selected for MS/MS analysis. The MS/MS raw data was analyzed by first converting into MS2 files, followed by database search using ProLuCID (Xu et al., 2006). The UWO241 protein database was generated based on our transcriptomics data supplemented with 37 common contaminants, and their reversed sequences as quality control system to restrain false positive discovery to 0.05. Differentially expressed proteins were analyzed using PatternLab for Proteomics (Carvalho et al., 2008). The proteomics results have been deposited to the MassIVE repository with the identifier MSV000084382. For identifying protein components in the supercomplex, the complex was harvested and 30 μg of total protein was processed similarly as described above to get the digested peptides. Different from the whole cell proteomics, the processed peptides were directly loaded onto a capillary C18

40 column without fractionation, and further analyzed in a Thermo LTQ Orbitrap XL mass spectrometer. The full mass spectra were recorded in the range of 350-1800 m/z with the resolution of 30,000. The top 12 peaks of each scan were selected for MS/MS analysis. The data analysis was conducted similarly as described above. Homologues of Lhca proteins were identified from a transcriptome previously generated from UWO241 (Raymond et al., 2009; NCBI BioProject No. PRJNA575885).

2.2.11 Gas Chromatography - Mass Spectrometry.

For determination of the primary metabolome UWO241 were grown in four biological replicate cultures as described above. Algal cells were harvested by centrifugation (6,000g, 5 min, 4°C) and washed once with fresh media. The supernatant was decanted, and the algal cells were flash frozen in liquid nitrogen and stored at -80°C. The metabolite extraction protocol was adapted from (Fiehn et al., 2008). In brief, metabolites were extracted from 20 mg of frozen tissue in 1 ml cold extraction buffer (methanol: chloroform: dH2O; 5:2:2). The samples were homogenized using glass beads (500 μm i.d.) in a Geno/Grinder 2010 instrument (SpexSamplePrep, Metuchen, NJ, USA), followed by centrifugation (14,000g, 2 min, 4°C). Samples were further processed and derivatized for GC-TOF mass spectrometry as described (Lee and Fiehn, 2008). GC-MS measurements were carried out on an Agilent 6890 gas chromatograph (Agilent, Santa Clara, CA, USA), controlled by a Leco ChromaTOF software v 2.32 (Leco, St. Joseph, MI, USA). Separation was performed on a Rtx-5Sil MS column (30m x 0.25mm x 0.25μm) with an additional 10 m empty guard column (Restek, Bellefonte, PA, USA) using helium as a carrier (1 ml/min flow rate). The oven temperature was held constant at 50°C for 1 min, the ramped at 20°/min to 330°C at which it was held constant for 5 min. A Leco Pegasus IV mass spectrometer (Leco, St. Joseph, MI, USA) was operated in electron impact (EI) mode at -70 eV ionization energy with unit mass resolution at 17 spectra/s with a scan range of 80-500 Da. The transfer line temperature between gas chromatograph and mass spectrometer was set to 280°C. Ionization Electron impact ionization at 70V was employed with an ion source temperature of 250°C. Mass spectra were processed using BinBase, an application system for deconvoluting and annotating mass spectral data, and analyzed as described in (Fiehn et al., 2005). Metabolites were identified based on their mass spectral characteristics and GC retention times, by comparison with compounds in a plant and algae reference library (West Coast Metabolomics Center, UC

Davis, CA, USA). Peak heights for the quantification ion at the specific retention index corresponding to each metabolite were normalized by the sum of peak heights in the sample. Normalized data were processed by cube root transformation followed by range scaling (van den Berg et al., 2006). Statistical analyses were performed by the Metaboanalyst 4.0 software suite (Chong et al., 2018), and included principal component analysis (PCA), t-test, heatmap and clustering analysis using Ward’s linkage for clustering and Pearson’s correlation as a measure of dissimilarity.

2.3 RESULTS

2.3.1 UWO241 is adapted to low temperature and high salt.

UWO241 was isolated from the deep photic zone (17 m sampling depth) of the hypersaline, perennially ice-covered lake (Lake Bonney, McMurdo Dry Valleys, Victoria Land) (Neale and Priscu, 1990; Neale and Priscu, 1995). As a consequence of more than two decades of study, this photopsychrophile has emerged as a model for photosynthetic adaptation to permanent low temperatures (Morgan-Kiss et al., 2006; Dolhi et al., 2013; Cvetkovska et al., 2017). In addition to psychrophily, UWO241 exhibits remarkable growth and photosynthetic performance under high salt (0.7 M NaCl, Appendix Fig. 2.1; Morgan et al., 1998; Pocock et al., 2011).

2.3.2 UWO241 possesses constitutively high rates of CEF.

PSI activity was monitored by far red (FR) light inducible P700 photooxidation (Fig. 2.1). Following a rise in absorbance at 820 (A820), reflecting FR-induced P700 oxidation, we compared rates of P700 re-reduction in the dark in LS cultures of C. reinhardtii as well as LS- and HS-grown cells of UWO241 (Fig. 2.1). Since FR preferentially excites PSI and not PSII, reduction of P700 following FR exposure is mainly due to alternative electron donors (Ivanov et al., 1998). UWO241 grown in standard LS growth medium exhibited a significantly shorter re- + red reduction time for P700 (t½ ) compared with LS-grown C. reinhardtii (p<0.05; Fig. 2.1). red Moreover, HS-grown UWO241 exhibited a 1.71-fold faster t½ compared with LS-grown cultures (p<0.05; 170±55 vs. 290±41 ms respectively; Fig. 2.1). The P700 absorbance kinetics also reveal that the oxidation of P700 in UWO241 is less complete in HS condition (Appendix Fig. 2.8). These data indicate that relative to the model C. reinhardtii, UWO241 exhibits a high

42 capacity for PSI-driven CEF, which is further enhanced during acclimation to long-term high salinity stress. Higher rates of CEF in HS-grown UWO241 were also confirmed by electrochromic shift (ECS) kinetics which estimates transthylakoid proton flux driven by light-dependent photosynthesis (Fig. 2.2, Appendix Fig. 2.2). The ECS signal was measured by the change in absorbance of thylakoid pigments at 520 nm during application of light dark interval (DIRK)

(Baker et al., 2007). The total amplitude of ECS signal (ECSt) was used to estimate the total proton motive force (pmf) across thylakoid membranes (Kramer et al., 2003). UWO241 grown in

HS exhibited 6 to 7.5 fold higher ECSt than that of LS-grown cells under all light intensities (Fig. 2.2 A), suggesting HS-grown cells generate higher pmf than LS-grown cells at the same light intensity. High pmf can be caused by either increased proton flux from LEF or CEF, reduced proton efflux, or decreased ATP synthase activity (Kanazawa and Kramer, 2002; Livingston et al., 2010; Carrillo et al., 2016). To verify which process(es) were contributing to high pmf in HS- + + grown cells, proton conductance (gH ) and fluxes through ATP synthase activity (νH ) were analyzed. The inverse of the lifetime of the rapid decay of ECS (ɡH+) represents proton permeability or conductivity of the thylakoid membrane and is largely dependent on the activity + of ATP synthesis (Appendix Fig. 2.2 D; Baker et al., 2007). The ɡH of HS-grown cells was ~50 + to 60% of that of LS-grown cells (Fig. 2.2 B); however, the proton flux rate (νH ) showed that the amount of ATP produced was still higher in HS-grown cells (Fig. 2.2 C). The relationship between νH+ and LEF can be used to estimate proton contribution from CEF (Baker et al., + 2007). In the linear plots of νH versus LEF, the slope of the HS-grown cells was higher than that of LS-grown cells (Fig. 2.2 D), indicating that CEF contributes significantly to the total proton + effluxes (νH ) in HS-grown cells. In close agreement with our P700 findings, UWO241-HS exhibited higher rates of CEF compared to UWO241-LS (Figs 2.1 and 2.2 D). Last, HS-grown cells exhibited downregulation of PSII and increased capacity for NPQ (Appendix Fig. 2.2 A and B), while PSI photochemical yield (Y(PSI)) was higher in HS- vs. LS-grown cells due to reduced PSI acceptor side limitation (Y(NA)) (Appendix Fig. 2.3).

2.3.3 Isolation of a PSI-supercomplex in UWO241.

Formation of PSI-supercomplexes have been shown to be essential for induction of CEF in plants and algae (DalCorso et al., 2008; Iwai et al., 2010). An earlier report showed that high

43 salinity-acclimated cultures of UWO241 form a PSI supercomplex (UWO241-SC); however, the yield of the UWO241-SC from fractionated thylakoids was relatively low and only a few proteins were identified (Szyszka-Mroz et al., 2015). In agreement with this report, the sucrose gradient from thylakoids isolated from LS-UWO241 had 3 distinct bands corresponding to major LHCII (Band 1), PSII core complex (Band 2) and PSI-LHCI (Band 3, Fig. 2.3 A). In contrast, UWO241-HS thylakoids lacked a distinct PSI-LHCI band, but exhibited several heavier bands, including the UWO241-SC band (Band 4; Fig. 2.3 B). We significantly improved recovery of the UWO241-SC by solubilizing thylakoids with the detergent a-DDM rather than β-DDM, which was used by other groups (Fig. 2.3 B). As a control, we compared the banding pattern of C. reinhardtii thylakoids when cells were exposed to the treatment typically used by others induce SC formation (i.e. State 1 vs. State 2 conditions, Iwai et al., 2010; Takahashi et al. 2013), (Fig. 2.3 C, D). In contrast with UWO241, band 3 (PSI) was more prominent while the SC band (band 4) was diffuse in State 2-treated C. reinhardtii cells (Fig. 2.3 D). Low-temperature fluorescence spectra were analyzed for the four bands extracted from the sucrose density gradients shown in Fig. 2.3 B and D (i.e. HS-UWO241 and State 2-treated C. reinhardtii, respectively). In C. reinhardtii, Band 1 exhibited a major emission peak at 680 nm (Fig. 2.4 C), corresponding to fluorescence from LHCII (Krause and Weis, 1991). Band 2 exhibited emission peak at 685 nm, consistent with PSII core (Fig. 2.4 A). Band 3 exhibited a peak at 685 nm and a strong peak at 715 nm; the latter consistent with PSI-LHCI (Fig. 2.4 A). However, Band 4 exhibited a strong fluorescence peak at 680 nm and a minor peak at 715 nm (Fig. 2.4 A). Fractionated thylakoids from HS-grown UWO241 exhibited emission spectra for Band 1 and Band 2 which were comparable with that from C. reinhardtii (Fig. 2.4 B). In contrast, both Band 3 and Band 4 (PSI and SC bands, respectively) exhibited highly reduced or a lack fluorescence associated with PSI. However, we confirmed the presence of the PSI reaction center protein, PsaA, in the UWO241-SC by immunoblotting (Fig. 2.4 C).

2.3.4 Protein composition of the supercomplex.

Protein components of the UWO241-SC were analyzed using LC-MS/MS. We identified a total of 39 proteins in the isolated band 4. The most abundant proteins in the supercomplex were proteins of the PSI reaction center and cytochrome b6f. In total we identified seven out of 13

44 subunits of PSI reaction center (Table 2.1; Appendix Table 2.1). Only two LHCI subunits, Lhca3 and Lhca5, and one LHCII minor subunit, CP29 were associated with the UWO241-SC. The Calcium sensing receptor (CAS) was identified as the third most abundant protein in the UWO241-SC. We also identified four subunits of ATP synthase in the UWO241-SC (α, β, γ, δ). Previously identified FtsH and PsbP protein were also detected. Bands 3 and 4 contained many PSI core proteins but lacked almost all Lhca proteins (Appendix Table 2.1). These results agreed with an earlier report that failed to detect most Lhca proteins in UWO241 thylakoids (Morgan et al., 1998). To determine whether the absence of Lhcas in the UWO241 proteome was due to a loss of Lhca genes, we searched a UWO241 transcriptome generated from a culture grown under low temperature/high salinity (Raymond et al., 2009). We identified 9 Lhca homologues which were transcribed under high salinity, suggesting that all or most of the LHCI genes are expressed in UWO241 (Appendix Fig. 2.4).

2.3.5 Whole cell proteome analysis.

We wondered whether constitutively high CEF could be linked to changes in downstream metabolism in UWO241. To address this question, whole-cell proteomes extracted from cultures of UWO241-HS and -LS grown under optimal temperature/light conditions were compared. Overall 98 proteins from various functional categories were identified as significantly affected in the two treatments, out of which 46 were upregulated and 62 downregulated in HS-acclimated cells. Proteins associated with photosynthesis (18%) and translational machinery (18%) were most affected in the treatment followed by primary and secondary metabolism (16%) (Appendix Fig. 2.5).

Photosynthesis: Differentially regulated proteins participating in photosynthesis were identified in UWO241-HS. Three photosystem reaction center proteins, PsaB and PsaN of PSI, and D2 protein of PSII, as well as extrinsic proteins of the water oxidizing complex, PsbO and PsbQ, were downregulated in UWO241-HS (Figs. 2.5 and 2.6 A; Appendix Table 2.2). One protein of chloroplastic ATP synthase, the epsilon subunit, was upregulated (3.8-fold) in UWO241-HS. Last, both FtsH proteins that were detected in the UWO241-SC, FtsH1 and FtsH2, were upregulated in UWO241-HS (Table 2.1; Appendix Table 2.2).

Several proteins associated with the CBB were significantly upregulated in UWO241-HS cells (Figs. 2.5 and 2.6A; Appendix Table 2.2). The RubisCO large subunit was upregulated in UWO241-HS, along with two chaperone proteins involved in RubisCO assembly, RuBA and RuBB. A class 2 fructose-1,6 –bisphosphatase (FBPase) was the third highest upregulated protein (5-fold), and fructose bisphosphate aldolase and transketolase were also upregulated in UWO241-HS (Fig. 2.6 A).

Metabolism: The enzyme glycerol-3-phosphate dehydrogenase (G3PDH), involved in glycerol- 3-phosphate biosynthesis, was the highest upregulated enzyme in UWO241-HS cultures (6-fold) (Fig. 2.6 B; Appendix Table 2.2). Fructose bis-phosphate aldolase, involved in gluconeogenesis and feeding into starch synthesis, as well as glucose-1-phosphate adenylyltransferase and starch synthase 1 (the last enzyme in starch synthesis pathway) were also upregulated in UWO241-HS (Fig. 2.6 B; Appendix Table 2.2). Two key enzymes from the shikimate pathway were significantly upregulated UWO241-HS: (i) the first enzyme in the pathway, DAHP (3-Deoxy-D-arabinoheptulosonate 7-phosphate) synthase (2.3-fold), and (ii) the last enzyme in the pathway, chorismate synthase (1.9-fold) (Fig. 2.6 B; Appendix Table 2.2). We found that indole-3-glycerol phosphate synthase (IGP synthase) was upregulated significantly in the HS conditions.

2.3.6. Primary metabolome analysis.

We detected a total of 771 unique metabolic signatures, 179 of which were positively identified based on their mass spectra and retention times (Kind et al., 2009). A heat map of all measured (identified and unidentified) metabolites showing the relative changes in primary metabolite abundances indicated clustering and a discrete population of metabolites that accumulate at high levels in HS-grown cultures when compared to LS-grown cells (Appendix Fig. 2.7). Overall, 59 metabolites (32%) were significantly different (p<0.01), out of which 9 were present in higher abundance and 50 were present in lower abundance in HS- vs. LS-grown UWO241 cultures (Appendix Tables 2.4 and 2.5). We observed high levels of glycerol in the primary metabolome of HS-grown UWO241 (8.7 FC), and high accumulation of the compatible solutes, sucrose (18.2 FC) and proline (27.1 FC) (Appendix Table 2.4). We also observed a high accumulation of phytol (12.6 FC),

suggesting chlorophyll degradation. HS-grown UWO241 cultures exhibited decreased amounts of 17 amino acids and compounds associated with amino acid metabolism (Appendix Table 2.5). Most notable metabolites from this class were lysine (29.4 FC) and ornithine (14.0 FC), which could signify a shift in amino acid metabolism to proline during exposure to high salinities. We also observed lower levels of the amino acid tryptophan (2.8 FC) in HS grown cultures. Metabolites involved in purine and pyrimidine metabolism were present in lower amounts in UWO241 exposed to high salinity, suggesting that these cells have shifted their metabolism from maintenance of the cell cycle and nucleic acid synthesis to producing osmoprotectants and compatible solutes. We also observed a reduction of 3-phosphoglycerate (3-PGA; 2.9 FC) in HS- grown cultures.

2.4 DISCUSSION

Our study shows that UWO241 maintains remarkable growth and photosynthesis under the combined stress of low temperature and high salt. This ability differs markedly from other model plants and algae that typically display significant downregulation of photosynthesis and growth when exposed to environmental stress, mainly as a consequence of bottlenecks in carbon fixation capacity (Hüner et al., 1998; Hüner et al., 2003; Ensminger et al., 2006; Hüner et al., 2016). Previous research has thoroughly described adaptive strategies for survival under permanent low temperatures, while survival under hypersalinity has received less consideration (Morgan et al., 1998; Morgan-Kiss et al., 2002a; Szyszka et al., 2007; Possmayer et al., 2011). One of the more distinct photosynthetic characteristics of UWO241 is the presence of a strong capacity for PSI-driven CEF (Morgan-Kiss et al., 2002b; Szyszka-Mroz et al., 2015; Cook et al., 2019). We validated that CEF rates are high in HS-grown cultures using the ECS signal, which was purported to mitigate problems with using P700 absorbance changes for CEF estimates (Lucker and Kramer, 2013). While CEF appears to be essential in plants and algae for balancing the ATP/NADPH ratio and protecting both PSI and PSII from photo-oxidative damage (Munekage et al., 2004; Joliot and Johnson, 2011; Huang et al., 2012), most studies report that CEF is part of short-term stress acclimation. Our work here as well as others suggests a larger role for CEF during long-term adaptation under permanent environmental stress (Morgan-Kiss et al., 2002b; Szyszka-Mroz et al., 2015; Cook et al., 2019a).

Constitutively high rates of CEF in UWO241 are associated with a general reorganization of PSI and PSII complexes (Szyszka-Mroz et al., 2019) and the formation of a Cyt bf-PSI supercomplex (Szyszka-Mroz et al., 2015). In this current study, following optimization of thylakoid protein complex solubilization by substituting β-DDM with a-DDM, the vast majority of PSI shifts from free PSI in the LS-grown cultures to association with the UWO241-SC in the HS-grown cultures. PSI supercomplexes have been described in several plant and algal species (Iwai et al., 2010; Li et al., 2018; Steinbeck et al., 2018). The UWO241-SC is distinct from that of C. reinhardtii because: i) its assembly is independent of short-term exposure to dark anaerobic conditions or other state transition-inducing treatments (Fig. 2.3), ii) the vast majority of PSI in UWO241 is associated with the UWO241-SC (Fig. 2.3), and iii) isolated UWO241-SC and PSI bands as well as whole cells lack typical PSI fluorescence emission at 77K, despite the presence of several PSI core proteins (Fig. 2.4; Table 2.1; Appendix Table 2.1; Morgan et al., 1998; Morgan-Kiss et al., 2002a, b; Cook et al., 2019). Thus, CEF combined with significant structural and functional changes to PSI are major targets for long-term stress acclimation in UWO241 (Fig. 2.7 A). While the UWO241-SC contains most of the PSI core proteins, both the UWO241-SC and PSI bands, as well as whole cell proteomes isolated from LS and HS conditions lacked homologues for most LHCI proteins (Table 2.1; Appendix Table 2.1). This agrees with an earlier study which was unable to detect most of the LHCI polypeptides by immunoblotting in UWO241 thylakoids (Morgan et al., 1998). Cook et al. (2019) also reported that the absence of LHCI proteins in UWO241 was not associated with adaptation to chronic iron deficiency, an additional stress experienced by natural communities of this alga. Transcriptomic analyses detected nine Lhca homologues in UWO241 grown under low temperature/high salinity (Appendix Fig. 2.4). Therefore, it appears that while most of the Lhca genes are encoded for and transcribed, few of the LHCI proteins are produced under the growth conditions tested thus far. These results fit well with numerous unsuccessful attempts to elicit typical 77K PSI long wavelength fluorescence emission in UWO241 (Morgan et al., 1998; Morgan-Kiss et al., 2002a; Morgan-Kiss et al., 2002b; 2005; Szyszka et al., 2007; Cook et al., 2019). It also explains the differences in the 77K emission spectra of the UWO241-SC and PSI bands between UWO241 and C. reinhardtii (Fig. 2.4). Last, a recent study reported that UWO241 transfers light energy from PSII to PSI via constitutive energy spillover through an undescribed mechanism (Szyska-

Mroz et al., 2019). Thus, UWO241 favors downregulated LHCI and constitutive energy spillover in response to its extreme habitat, most likely the natural light environment of extreme shade enriched in blue wavelengths (Neale and Priscu, 1995). The direct product of CEF is extra transthylakoid proton motive force at the expense of NADPH production (Lucker and Kramer, 2013; Dumas et al., 2016; Yamori et al., 2016a), with the consequences of CEF impacting either cellular energy production or photoprotection. CEF- dependent formation of ΔpH protects PSII by activating the energy-dependent quenching (qE), a major process for dissipation of excess light energy in PSII (Yamori et al., 2016). Alternatively, CEF-generated pmf can be used for production of additional ATP for high-energy consuming processes including protein synthesis, transport processes, ion homeostasis (He et al., 2015), CO2 concentrating mechanisms (Horváth et al., 2000; Lucker and Kramer, 2013), or production of secondary metabolites (Murthy et al., 2014). Last, CEF prevents PSI photoinhibition by downregulating LEF and alleviating over-reduction of the acceptor side of PSI, thereby preventing ROS-induced PSI damage (Munekage et al., 2008; Shimakawa et al., 2016; Chaux et al., 2017; Huang et al., 2017). A strong constitutive CEF mechanism in UWO241 could be beneficial for one or most of the above purposes. First, HS-grown cultures possess a higher capacity for NPQ (Appendix Fig. 2.2), supporting a role for CEF in constitutive photoprotection ability. High CEF rates also correlate with a higher Y(PSI) and a lower PSI acceptor side limitation in HS-grown cultures (Appendix Fig. 2.3), suggesting enhanced PSI photoprotection in UWO241-HS cells. UWO241-HS cells exhibited significantly higher levels of cellular ATP compared with that of UWO241-LS cells (Appendix Fig. 2.9). In addition, ATP synthase subunits were associated with the UWO241-SC (Table 2.1), suggesting CEF contributes extra ATP in UWO241. HS-grown

cultures exhibited significantly higher ECSt and νH+ compared to LS-grown cultures, suggesting a high flux of protons through the chloroplastic ATP synthase in spite of slow activity of ATP synthase (compare Fig. 2.2 A and C with Fig. 2.2 B). Slower activity of ATP synthase could be overcome by higher abundance of ATP synthase subunits in the HS-grown UWO241 which is reported here (Figs. 2.5 and 2.6) and in an earlier report (Morgan et al., 1998). Recently it was shown that in a salt-tolerant soybean, increased CEF contributes to excess ATP that is used to drive import of Na+ in the vacuole (He et al., 2015). Taken together, constitutively high rates of CEF in UWO241 are likely to provide dual benefits, that of constitutive photoprotection of both

PSI and PSII and extra ATP to support downstream processes important for low temperature and/or high salinity adaptation (Fig. 2.7 A). Comparison of whole cell proteomes and metabolomes revealed significant shifts in primary metabolism in LS- and HS-grown UWO241. First, HS-grown cultures have a strong carbon fixation potential. Key enzymes within the CBB cycle are upregulated under HS, including large subunit of RuBisCO (LSU, EC 4.1.39) and a chaperone complex involved in RuBisCO assembly (RuBA and RuBB proteins). Several enzymes important in regeneration of ribulose-1,5-bisphosphate (RuBP), including fructose-1,6-bisphosphatase (FBPase, EC 3.1.3.11), fructose-bisphophate aldolase (EC 4.1.2.13), transketolase (EC 2.2.1.1), ribose-5 phosphate isomerase (EC 5.2.1.6), and phosphoribulokinase (PRK, EC 2.7.1.19), are also higher in HS- grown cells (Fig. 2.7 B, represented by thick arrows). Overexpression of key bottleneck CBB enzymes such as FBPase and SBPase enhances carbon fixation and RuBP regeneration (Lefebvre et al., 2005; Tamoi et al., 2006), while also supporting improved photosynthesis during stress (Driever et al., 2017). Low levels of 3-PGA, the direct biproduct of RubisCO activity also suggests strong carbon sinks for fixed CO2 in HS-grown cultures (Appendix Table 2.5). Last, overproduction of these key CBB enzymes is supported by a robust protein translation ability, as several ribosomal proteins are also overexpressed in HS-grown cells (Appendix Table 2.2). Enhanced CBB pathway activity would support robust photosynthetic activity and growth in UWO241. However, proteomic evidence revealed other potential carbon sinks, including carbon storage in the form of starch (Appendix Table S2). Two key enzymes of starch synthesis were upregulated under high salinity, G1P adenylyltransferase (AGPase; EC 2.7.7.27) and starch synthase (EC 2.4.1.242). AGPase catalyzes the formation of glucose-1-phosphate to ADP- glucose and consumes 1 ATP. ADP-glucose serves as substrate for starch synthase to extend the glucosyl chain in starch. In contrast with these findings, plant and algal fitness and survival under low temperatures or high salinity is associated with starch degradation (reviewed in Thalmann et al., 2016). Under abiotic stress starch is remobilized into sugars and other metabolites to provide carbon and energy when photosynthesis is compromised. In contrast with cold- or salt-sensitive plants and algae, the photosynthetic apparatus of UWO241 is remodeled to support photosynthesis under continuous low temperatures and high salinity (Morgan et al., 1998; Szyszka et al., 2007; Pocock et al., 2011). Thus, accumulation of starch in UWO241 may act as a strong carbon sink to support high rates of carbon fixation (Fig. 2.7 B). Starch stored in

the chloroplast is also transitory, and is often rapidly turned over (Thalmann et al., 2016). Starch content was comparable between LS- and HS-grown UWO241 cultures (Appendix Fig. 2.6). Thus, transiently stored starch could be an additional adaptive strategy in UWO241, acting as an energy and carbon buffer which can be rapidly mobilized when needed. This theory is supported by other publications that reported accumulation of starch under cold or salinity stress (Siaut et al., 2011; Wang et al., 2013), suggesting that transitory starch synthesis and mobilization may be important during stress acclimation. The shikimate pathway is an essential link between primary and secondary metabolism for producing precursors for aromatic amino acids (tryptophan, phenylalanine, tyrosine) as well as many other aromatic metabolites such as, indole compounds, alkaloids, lignin and flavonoids (Maeda and Dudareva, 2012). It is a high carbon flux pathway, accounting for approx. 30% of all

fixed CO2 in an organism (Tohge et al., 2013). Various biotic and abiotic stresses upregulate genes of the shikimate pathway as well as downstream biosynthesis pathways which use the main product of the shikimate pathway, chorismate, as a substrate (Tzin and Galili, 2010). HS- grown UWO241 exhibited upregulation of two key Shikimate enzymes, 3-deoxy-D-arabinoheptulosonate 7-phosphate synthase (DHAP synthase; EC 2.5.1.54) and chorismate synthase (EC 4.2.3.5). The substrates for DAHP synthase are erythrose-4-phosphate (E4P) and PEP. E4P is a product of the CBB cycle enzyme transketolase. Upregulation of transketolase in HS-grown cells indicates that E4P would be supplied in high levels to support high flux through the Shikimate pathway. On the other hand, supply of PEP for chorismate synthesis is likely to come from glycolysis, as indicated by the higher level of expression of G3P dehydrogenase under the HS condition (Appendix Table 2.2). Thus, the CBB cycle and glycolysis are likely to be coordinated in order to provide substrates to support high flux through the shikimate pathway. Last, there is evidence linking CEF with the Shikimate pathway. One recent study involving a CEF mutant of Arabidopsis thaliana (lacking pgr5 protein) showed that the levels of shikimate metabolites were significantly reduced in the CEF mutant as compared to wild type, suggesting a link between CEF and chorismate synthesis (Florez-Sarasa et al., 2016). Acclimation to a variety of stresses in plants and algae often involves upregulation of heat shock proteins, stress metabolites, as well as signaling molecules such as plant hormones and signal transduction pathways (eg: Ca 2+) (Montero-Barrientos et al., 2010; Suzuki et al., 2016). We provide evidence in the proteome of UWO241 that a biosynthetic enzyme of the

tryptophan pathway, indole-3-glycerol phosphate synthase (IGPS, EC 4.1.1.48) is highly expressed under high salt; however, the metabolome data showed that L-Trp levels were reduced in the HS-grown cultures (Appendix Tables 2.2 and 2.5). L-Trp is also a major substrate for production of the phytohormone, indole-3-acetic acid (IAA), and the product of IGPS, indol-3- glycerol phosphate, is a branch point between L-Trp synthesis and a L-Trp independent IAA synthesis pathway (Ouyang et al., 2000). IAA and several other phytohomormones have been detected in a few cyanobacteria and algal species; however, their putative function is largely based on exogenously added plant phytohormones to algal cultures (Lu and Xu, 2015). Exogenously added IAA stimulates carbon fixation and growth and enhances stress tolerance in algae (Lu and Xu, 2015). Last, IAA production increases Ca2+ levels in plants during acclimation to abiotic stress (Vanneste and Friml, 2013). Ca2+ signaling has been linked to both CEF and assembly of PSI supercomplexes (Terashima et al., 2012). Indeed, the Calcium sensing receptor (CAS) was an abundant protein associated with the UWO241-SC (Table 2.1). More work will be needed to ascertain whether IAA and Ca2+ play roles in CEF and assembly of the UWO241-SC. Despite more than 2 decades of study on UWO241, there are many questions that remain unanswered. Recent papers have added to the breadth of knowledge on the photosynthetic apparatus, including a cold-adapted ferredoxin isoform (Cvetkovska et al., 2018) and a thylakoid kinase exhibiting temperature-dependent phosphorylation patterns (Szyska-Mroz et al., 2019). Here we extend our understanding of long-term photosynthetic adaptation to permanent cold and hypersalinity by proposing a model for sustained PSI-CEF that supports a robust CBB pathway and a regular growth rate (Fig. 2.7). Under permanent environmental stress, CEF supplies constitutive photoprotection of PSI and PSII while also producing extra ATP for downstream metabolism (Fig. 2.7 A). The restructured photosynthetic apparatus is accompanied by major rewiring of central metabolism to provide a strong carbon fixation potential which is used in part to produce stored carbon and drive high carbon flux pathways (Fig. 2.7 B). Algae adapted to multiple stressors such as low temperatures combined with high salinity are robust fixers of CO2, providing new genetic targets for improving crop stress resistance and previously unconsidered sources of natural carbon sinks.

Table 2.1. The UWO241 supercomplex contains subunits of PSI, Cytb6f, ATP synthase, as well as several antenna proteins. Proteomic analysis of Band 4 isolated from cultures acclimated to high salinity (700 mM NaCl). NSAF (percent normalized spectral abundance factor) for selected proteins are shown.

Protein NSAF (%) UniProtKB Organism PSI PsaD 4.53 Q39615 C. reinhardtii PsaE 4.24 P12356 C.reinhardtii PsaF 3.12 P12356 C.reinhardtii PsaH 1.84 P13352 C. reinhardtii PsaK 6.13 P14225 C. reinhardtii PsaL 2.43 Q39654 C. sativus LHCI & LHCII Antenna Lhca3 1.35 Q9SY97 A. thaliana Lhca5 2.82 Q9C639 A. thaliana CP29 2.25 Q93WD2 C. reinhardtii LHCII type I 2.22 P20866 P. patens CB2 9.46 P14273 C.reinhardtii Cyt b6f PetA 5.35 P23577 C. reinhardtii PetB 2.33 Q00471 C. reinhardtii PetC 1.15 P49728 C.reinhardtii ATP Synthase AtpA 2.45 P26526 C. reinhardtii AtpB 2.07 P06541 C. reinhardtii Other CAS 4.65 Q9FN48 A. thaliana FtsH1 0.49 Q5Z974 O. sativa FtsH2 1.26 O80860 A. thaliana PsbP 2.52 P11471 C. reinhardtii

a C. reinhardtii t1/2 = 509 ± 46 ms 1.00 b UWO241 LS t1/2 = 290 ± 41 ms

) c UWO241 HS t1/2 = 170 ± 55 ms 820 A

0.75

0.50

reduction ( + 700

P 0.25

0.00 0 1000 2000 3000 Time (ms)

Figure 2.1. P700 re-reduction kinetics of photosystem I in C. sp. UWO241 compared to the model mesophile C.reinhardtii. C. reinhardtii was grown in low salt BBM medium at 20oC/100 μmol m-2s-1. UWO241 was grown in either BBM (low salt, LS) or BBM+700 mM NaCl (High -2 1 salt, HS), and 8°C/50 μmol m s- . t1/2, half-time for P700 re-reduction. Values labelled with different lower-case letters indicate statistical difference (n=6-9; p< 0.05).

0.0012 45 A 40 B 0.001 35 0.0008 30 ) t 1 - 25 0.0006 ■ UWO241 LS (s

ECS H+ 20 ● UWO241 HS g 0.0004 15 10 0.0002 5 0 0 0 100 200 300 400 500 600 0 100 200 300 400 500 600 Light intensity (µmol photons m-2 s-1) Light intensity (µmol photons m-2 s-1)

0.00002 0.000016 C 0.000014 D 0.000016 0.000012

0.000012 0.00001

H+ H+ 0.000008 ν ν 0.000008 0.000006

0.000004 0.000004 0.000002

0 0 0 100 200 300 400 500 600 0 10 20 30 40 50 Light intensity (µmol photons m-2 s-1) LEF (µmol photons m-2 s-1)

Figure 2.2. Photosynthetic properties of C. sp. UWO241 under various light intensities using

Electrochomic shift. The ECSt (total amplitude of ECS, proportional to proton motive force, A), gH+ (proton conductance, reflecting the ATP synthesis activities, B) and vH+ (proton flux rates, C) of C. sp. UWO241 grown in low-salt (black, squares) and high-salt (grey, circles) were measured from dark interval relaxation kinetics (DIRK). The relationship between vH+ and LEF (measured by chlorophyll fluorescence, see Appendix Figure 2.2) were assessed in C. sp. UWO241 (D).

C. sp. UWO241 C. reinhardtii LS HS State 1 State 2

1 1 1 1 2 2 3 2 2 3 3 3

4 4 AB CD

Figure 2.3. Supercomplex formation in C. sp. UWO241 vs. C. reinhardtii. Fractionation of major thylakoid chlorophyll-protein complexes from C. sp. UWO241 by sucrose density ultracentrifugation from low salt (LS) - and high salt (HS)- grown cultures (A, B). Fractionation of major thylakoid chlorophyll-protein complexes from C. reinhardtii exposed to State 1 and State 2 conditions (C, D). Cultures of C. reinhardtii were grown under control conditions (20°C/100 umol) and either dark adapted for 10 minutes (State 1, C) or incubated under anaerobic conditions for 30 minutes (State 2, D).

A Band 1 Band 2 685 715 1.00 680 Band 3 Band 4

0.75

0.50

0.25 FluorescenceYield (R. U.)

0.00 675 700 725 750

B Band 1 Band 2 1.00 683 Band 3 Band 4

0.75 C 75

0.50 50

715 0.25 FluorescenceYield (R.U.)

0.00 675 700 725 750 Emission Wavelength (nm)

Figure 2.4. 77K fluorescence emission spectra of Chl-protein complexes from C.reinhardtii and C. sp. UWO241. Low temperature Chl a fluorescence emission spectra of pigment-protein bands isolated from sucrose density gradients shown in Figure 2.3. Low temperature Chl a emission spectra of bands from C. reinhardtii State II conditions (A). Emission spectra of bands from high salt cultures of C. sp. UWO241 (B). Immunoblot of the UWO241-SC band with PsaA antibody (C). Spectra represent normalized data and the average of 3 scans.

Photosynthesis

CBB Cycle

Central Metabolism

Translation

Stress, membrane and transport proteins

Figure 2.5. Heat map of differentially regulated proteins in UWO241 under low (LS) and high salinity (HS) conditions. The normalized spectral abundance factor (NSAF) values are plotted for each replicate in the two conditions (n=3) using color based approach (green: low abundance, red: high abundance). The proteins are categorized into broad processes they belong to.

Photosynthesis Metabolism Primary 6 A PETC Storage Calvin Cycle B Secondary Amino Acid 4 5

2 0 0

-2 -5 Fold Change (HS vs LS) Fold Change (HS vs LS) -4

FTSH1FTSH2 RubisCO L Transketolase DAHP Synthase GDP-mannose Starch Synthase 1 Biotin Carboxylase Aconitate Hydratase Methionine synthase Phosphoribulokinase Pyruvate Carboxylase ATP synthase epsilon Malate DehydrogenaseChorismate SynthaseGlycine 1 dehydrogenase Argininosuccinate lyase Photosystem II D2 protein 15-cis-phytoene desaturase Ketol-acidSulfate reductoisomerase adenylyltransferase Fructose-1,6-bis phophatase Fructose bis-phosphate aldolase S-adenosylmethionine synthase Aldehyde-alcohol dehydrogenase Ribose-5-phosphate isomerase 3 Phosphomethylpyrimidine synthase Phosphoserine aminotransferase 1 Inorganic soluble pyrophosphatase 1 Oxygen-evolvingOxygen-evolving enhancer enhancer protein protein 1 3 Indole-3-glycerol phosphate synthase Aspartate-semialdehyde dehydrogenase Glutamate--glyoxylate24-methylenesterol aminotransferase 2 C-methyltransferase 2 Photosystem I reaction center subunit N Carbamoyl-phosphateNADH dehydrogenase synthase [ubiquinone] large chain 1 alpha RubisCo large subunit binding protein beta Glycerol-3-phosphateGlycerol-3-phosphate dehydrogenase dehydrogenase [NAD+] [NAD+] RubisCo large subunit binding proteinNAD(P)H-quinone alpha oxidoreductase subunit S Phospho-2-dehydro-3-deoxyheptonate aldolase1 Photosystem II stability/assembly factor HCF136 Photosystem I P700 chlorophyll a apoprotein A2 Glucose-1-phosphate adenylyltransferase large subunit

Figure 2.6. Changes in photosynthetic machinery and downstream metabolism in UWO241 associated with high salinity. Proteins participating in photosynthetic machinery (PETC: green, Calvin cycle: Blue) that are significantly affected in UWO241 under high salinity (A). Proteins participating in downstream metabolism that belong to primary (blue), storage (green), secondary (red) and amino acid biosynthesis (brown) metabolic pathways and are significantly affected in UWO241 under high salinity are shown (B). Proteins with fold change > 1.2 are shown (n=3). PETC: Photosynthetic electron transport chain.

A ADP + Pi ATP

NPQ spill-over

PSII Cytbf PSI CAS Lhca3 PC H O + 2 O2+H Cyt b6f-PSI supercomplex B

L-trp/IAA synthesis IGPS L-trp

Figure 2.7. A restructured photosynthetic apparatus supports rewiring of central metabolism in C. sp. UWO241. The photosynthetic apparatus of UWO241 is assembled to promoted high rates of

CEF which is sustained by formation of a PSI supercomplex (A). CEF supports photoprotection of

PSII and PSI and provides additional ATP for downstream metabolism. High ATP is consumed, in part, by the CBB cycle as well as an upregulated shikimate pathway and carbon storage pathways

(starch, glycerol) (B). Thick arrows represent significantly upregulated enzymes. Model is based on data presented here and other studies (Cook et al., 2019; Szyszka-Mroz et al., 2015; 2019)

2.5 REFERENCES

Avron M (1986) The osmotic components of halotolerant algae. Tren Biochem Sci 11: 5-6 Baker NR (2008) Chlorophyll fluorescence: a probe of photosynthesis in vivo. Ann Rev Plant Biol 59: 89-113 Baker NR, Harbinson J, Kramer DM (2007) Determining the limitations and regulation of photosynthetic energy transduction in leaves. Plant, Cell & Environ 30: 1107-1125 Ben-Amotz A, Avron M (1973) The role of glycerol in the osmotic regulation of the halophilic alga Dunaliella parva. Plant Physiol. 51: 875-878 Berens ML, Berry HM, Mine A, Argueso CT, Tsuda K (2017) Evolution of hormone signaling networks in plant defense. Ann Rev Phytopathol 55: 401-425 Brown A (1990) Microbial water stress physiology. Principles and perspectives. John Wiley & Sons Cardol P, Forti G, Finazzi G (2011) Regulation of electron transport in microalgae. Biochim et Biophys Acta 1807: 912-918 Carrillo LR, Froehlich JE, Cruz JA, Savage LJ, Kramer DM (2016) Multi-level regulation of the chloroplast ATP synthase: the chloroplast NADPH thioredoxin reductase C (NTRC) is required for redox modulation specifically under low irradiance. Plant J 87: 654-663 Carvalho PC, Fischer JS, Chen EI, Yates JR, Barbosa VC (2008) PatternLab for proteomics: a tool for differential shotgun proteomics. BMC Bioinform 9: 316 Chandra-Shekara A, Venugopal SC, Barman SR, Kachroo A, Kachroo P (2007) Plastidial fatty acid levels regulate resistance gene-dependent defense signaling in Arabidopsis. Proc Natl Acad Sci USA 104: 7277-7282 Chaux F, Johnson X, Auroy P, Beyly-Adriano A, Te I, Cuiné S, Peltier G (2017) PGRL1 and LHCSR3 compensate for each other in controlling photosynthesis and avoiding photosystem I photoinhibition during high light acclimation of Chlamydomonas cells. Molec Plant 10: 216-218 Chong J, Soufan O, Li C, Caraus I, Li S, Bourque G, Wishart DS, Xia J (2018) MetaboAnalyst 4.0: towards more transparent and integrative metabolomics analysis. Nucl Acids Res 46: W486-W494

Cook G, Teufel A, Kalra I, Li W, Wang X, Priscu J, Morgan-Kiss R (2019a) The Antarctic psychrophiles Chlamydomonas spp. UWO241 and ICE-MDV exhibit differential restructuring of photosystem I in response to iron. Photosynth. Res. 141: 209-228 Cvetkovska M, Hüner NP, Smith DR (2017) Chilling out: the evolution and diversification of psychrophilic algae with a focus on Chlamydomonadales. Polar Biol. 40: 1169-1184 Cvetkovska M, Szyszka‐Mroz B, Possmayer M, Pittock P, Lajoie G, Smith DR, Hüner NP (2018) Characterization of photosynthetic ferredoxin from the Antarctic alga Chlamydomonas sp. UWO 241 reveals novel features of cold adaptation. New Phytol 219: 588-604 DalCorso G, Pesaresi P, Masiero S, Aseeva E, Schünemann D, Finazzi G, Joliot P, Barbato R, Leister D (2008) A Complex Containing PGRL1 and PGR5 Is Involved in the Switch between Linear and Cyclic Electron Flow in Arabidopsis. Cell 132: 273-285 Darko E, Heydarizadeh P, Schoefs B, Sabzalian MR (2014) Photosynthesis under artificial light: the shift in primary and secondary metabolism. Phil Trans R Soc B 369: 20130243 Dolhi JM, Maxwell D, Morgan-Kiss RM (2013) The Antarctic Chlamydomonas raudensis: an emerging model for cold adaptation of photosynthesis. Extremophiles 17: 711-722 Driever SM, Simkin AJ, Alotaibi S, Fisk SJ, Madgwick PJ, Sparks CA, Jones HD, Lawson T, Parry MA, Raines CA (2017) Increased SBPase activity improves photosynthesis and grain yield in wheat grown in greenhouse conditions. Phil Trans R Soc Lond B. 372: 20160384 Driver T, Trivedi DK, McIntosh OA, Dean AP, Goodacre R, Pittman JK (2017) Two glycerol-3- phosphate dehydrogenases from Chlamydomonas have distinct roles in lipid metabolism. Plant Physiol. 174: 2083-2097

Dumas L, Chazaux M, Peltier G, Johnson X, Alric J (2016) Cytochrome b6f function and localization, phosphorylation state of thylakoid membrane proteins and consequences on cyclic electron flow. Photosynth. Res. 129: 307-320 Ensminger I, Busch F, Huner NPA (2006) Photostasis and cold acclimation: sensing low temperature through photosynthesis. Physiol. Plant. 126: 28-44 Fang L, Lin HX, Low CS, Wu MH, Chow Y, Lee YK (2012) Expression of the Chlamydomonas reinhardtii Sedoheptulose‐1, 7‐bisphosphatase in Dunaliella bardawil leads to enhanced photosynthesis and increased glycerol production. Plant Biotechnol J 10: 1129- 1135

Fiehn O, Wohlgemuth G, Scholz M (2005) Setup and annotation of metabolomic experiments by integrating biological and mass spectrometric metadata. In International workshop on data integration in the life sciences. Springer, pp 224-239 Fiehn O, Wohlgemuth G, Scholz M, Kind T, Lee DY, Lu Y, Moon S, Nikolau B (2008) Quality control for plant metabolomics: reporting MSI‐compliant studies. Plant J 53: 691-704 Florez-Sarasa I, Noguchi K, Araujo WL, Garcia-Nogales A, Fernie AR, Flexas J, Ribas-Carbo M (2016) Impaired Cyclic Electron Flow around Photosystem I Disturbs High-Light Respiratory Metabolism. Plant Physiol. 172: 2176-2189 Gimmler H, Möller EM (1981) Salinity‐dependent regulation of starch and glycerol metabolism in Dunaliella parva. Plant Cell Environ. 4: 367-375 Goyal A (2007a) Osmoregulation in Dunaliella, Part I: Effects of osmotic stress on photosynthesis, dark respiration and glycerol metabolism in Dunaliella tertiolecta and its salt-sensitive mutant (HL 25/8). Plant Physiology and Biochemistry 45: 696-704 Goyal A (2007b) Osmoregulation in Dunaliella, Part II: Photosynthesis and starch contribute carbon for glycerol synthesis during a salt stress in Dunaliella tertiolecta. Plant Physiol Biochem 45: 705-710 He Y, Fu J, Yu C, Wang X, Jiang Q, Hong J, Lu K, Xue G, Yan C, James A (2015) Increasing cyclic electron flow is related to Na+ sequestration into vacuoles for salt tolerance in soybean. J Exp Bot: erv392 Herrera-Valencia VA, Macario-Gonzalez LA, Casais-Molina ML, Beltran-Aguilar AG, Peraza- Echeverria S (2012) In silico cloning and characterization of the glycerol-3-phosphate dehydrogenase (GPDH) gene family in the green microalga Chlamydomonas reinhardtii. Curr Microbiol 64: 477-485 Horváth EM, Peter SO, Joët T, Rumeau D, Cournac L, Horváth GV, Kavanagh TA, Schäfer C, Peltier G, Medgyesy P (2000) Targeted inactivation of the plastid ndhB gene in tobacco results in an enhanced sensitivity of photosynthesis to moderate stomatal closure. Plant Physiol. 123: 1337-1350 Huang W, Yang S-J, Zhang S-B, Zhang J-L, Cao K-F (2012) Cyclic electron flow plays an important role in photoprotection for the resurrection plant Paraboea rufescens under drought stress. Planta 235: 819-828

Huang W, Zhang S-B, Xu J-C, Liu T (2017) Plasticity in roles of cyclic electron flow around photosystem I at contrasting temperatures in the chilling-sensitive plant Calotropis gigantea. Env Exp Bot 141: 145-153 Hüner NP, Dahal K, Bode R, Kurepin LV, Ivanov AG (2016) Photosynthetic acclimation, vernalization, crop productivity and ‘the grand design of photosynthesis’. J Plant Physiol 203: 29-43 Hüner NPA, Öquist G, Melis A (2003) Photostasis in plants, green algae and cyanobacteria: the role of light harvesting antenna complexes. In Advances in Photosynthesis and Respiration. Light Harvesting Antennas in Photosynthesis (Green BR, Parson WW, eds). Kluwer Academic Publishers, Dordrecht, Vol 13, pp 401-421 Hüner NPA, Öquist G, Sarhan F (1998) Energy balance and acclimation to light and cold. Trends Plant Sci 3: 224-230 Ishikawa N, Takabayashi A, Noguchi K, Tazoe Y, Yamamoto H, von Caemmerer S, Sato F, Endo T (2016) NDH-mediated cyclic electron flow around photosystem I is crucial for C4 photosynthesis. Plant Cell Physiol 57: 2020-2028 Ivanov AG, Morgan RM, Gray GR, Velitchkova MY, Huner NP (1998) Temperature/light dependent development of selective resistance to photoinhibition of photosystem I. FEBS Lett. 430: 288-292 Iwai M, Takizawa K, Tokutsu R, Okamuro A, Takahashi Y, Minagawa J (2010) Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis. Nature 464: 1210- 1213 Jeong J, Baek K, Kirst H, Melis A, Jin E (2017) Loss of CpSRP54 function leads to a truncated light-harvesting antenna size in Chlamydomonas reinhardtii. Biochim Biophys Acta 1858: 45-55 Joliot P, Johnson GN (2011) Regulation of cyclic and linear electron flow in higher plants. Proc Natl Acad Sci USA 108: 13317-13322 Kanazawa A, Kramer DM (2002) In vivo modulation of nonphotochemical exciton quenching (NPQ) by regulation of the chloroplast ATP synthase. Proc Natl Acad Sci USA 99: 12789- 12794

Kind T, Wohlgemuth G, Lee DY, Lu Y, Palazoglu M, Shahbaz S, Fiehn O (2009) FiehnLib: mass spectral and retention index libraries for metabolomics based on quadrupole and time-of- flight gas chromatography/mass spectrometry. Anal Chem 81: 10038-10048 Kramer DM, Cruz JA, Kanazawa A (2003) Balancing the central roles of the thylakoid proton gradient. Tren. Plant Sci. 8: 27-32 Kramer DM, Evans JR (2011) The importance of energy balance in improving photosynthetic productivity. Plant Physiol. 155: 70-78 Krause GH, Weis E (1991) Chlorophyll fluorescence and photosynthesis: the basics. Ann. Rev. Physiol. Plant Mol. Biol. 42: 313-349 Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680 Lee DY, Fiehn O (2008) High quality metabolomic data for Chlamydomonas reinhardtii. Plant Meth 4: 7 Lefebvre S, Lawson T, Fryer M, Zakhleniuk OV, Lloyd JC, Raines CA (2005) Increased sedoheptulose-1, 7-bisphosphatase activity in transgenic tobacco plants stimulates photosynthesis and growth from an early stage in development. Plant Physiol. 138: 451- 460 Li L, Aro E-M, Millar AH (2018) Mechanisms of photodamage and protein turnover in photoinhibition. Tren. Plant Sci. 23: 667-676 Livingston AK, Cruz JA, Kohzuma K, Dhingra A, Kramer DM (2010) An Arabidopsis mutant with high cyclic electron flow around Photosystem I (hcef) Involving the NADPH Dehydrogenase Complex. Plant Cell 22: 221-233 Lu Y, Xu J (2015) Phytohormones in microalgae: a new opportunity for microalgal biotechnology? Tren. Plant Sci. 20: 273-282 Lucker B, Kramer DM (2013) Regulation of cyclic electron flow in Chlamydomonas reinhardtii under fluctuating carbon availability. Photosynth. Res. 117: 449-459 Maeda H, Dudareva N (2012) The shikimate pathway and aromatic amino acid biosynthesis in plants. Annu Rev Plant Biol 63: 73-105 Magnuson A, Cardona T (2016) Thylakoid membrane function in heterocysts. Biochim Biophys Acta 1857: 309-319

Magnuson A, Krassen H, Stensjö K, Ho FM, Styring S (2011) Modeling Photosystem I with the alternative reaction center protein PsaB2 in the nitrogen fixing cyanobacterium Nostoc punctiforme. Biochim Biophys Acta 1807: 1152-1161 Minagawa J (2016) A Supercomplex of Cytochrome bf and Photosystem I for Cyclic Electron Flow. In Cytochrome Complexes: Evolution, Structures, Energy Transduction, and Signaling. Springer, pp 453-462 Montero-Barrientos M, Hermosa R, Cardoza RE, Gutierrez S, Nicolas C, Monte E (2010) Transgenic expression of the Trichoderma harzianum hsp70 gene increases Arabidopsis resistance to heat and other abiotic stresses. J Plant Physiol 167: 659-665 Morgan-Kiss R, Ivanov AG, Williams J, Mobashsher K, Hüner NP (2002a) Differential thermal effects on the energy distribution between photosystem II and photosystem I in thylakoid membranes of a psychrophilic and a mesophilic alga. Biochim Biophys Acta 1561: 251- 265 Morgan-Kiss RM, Ivanov A, Modla S, Cyzmmek K, Huner NPA, Priscu JC, Hanson TE (2008) Identity and phylogeny of a new psychrophillic eukaryotic green alga, Chlorella sp. strain BI isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles 12: 701- 711 Morgan-Kiss RM, Ivanov AG, Hüner NPA (2002b) The Antarctic psychrophile, Chlamydomonas subcaudata, is deficient in state I-state II transitions. Planta 214: 435-445 Morgan-Kiss RM, Ivanov AG, Pocock T, Król M, Gudynaite-Savitch L, Hüner NPA (2005) The Antarctic psychrophile, Chlamydomonas raudensis Ettl (UWO241) (CHLOROPHYCEAE, CHLOROPHYTA) exhibits a limited capacity to photoacclimate to red light. J Phycol 41: 791-800 Morgan-Kiss RM, Priscu JC, Pocock T, Gudynaite-Savitch L, Huner NP (2006) Adaptation and acclimation of photosynthetic microorganisms to permanently cold environments. Microbiol Molec Biol Rev 70: 222-252 Morgan RM, Ivanov AG, Priscu JC, Maxwell DP, Hüner NPA (1998) Structure and composition of the photochemical apparatus of the Antarctic green alga, Chlamydomonas subcaudata. Photosyn. Res. 56: 303-314

Munekage Y, Hashimoto M, Miyake C, Tomizawa K-I, Endo T, Tasaka M, Shikanai T (2004) Cyclic electron flow around photosystem I is essential for photosynthesis. Nature 429: 579- 582 Munekage YN, Genty B, Peltier G (2008) Effect of PGR5 impairment on photosynthesis and growth in Arabidopsis thaliana. Plant Cell Physiol 49: 1688-1698 Murthy HN, Lee E-J, Paek K-Y (2014) Production of secondary metabolites from cell and organ cultures: strategies and approaches for biomass improvement and metabolite accumulation. Plant Cell, Tissue and Organ Culture (PCTOC) 118: 1-16 Nawrocki W, Bailleul B, Picot D, Cardol P, Rappaport F, Wollman F-A, Joliot P (2019) The mechanism of cyclic electron flow. Biochimica et Biophysica Acta (BBA)-Bioenergetics Neale PJ, Priscu JC (1990) Structure and function of the photochemical apparatus in the phytoplankton of ice-covered Lake Bonney. Antarc. J. U.S. 25: 225-225 Neale PJ, Priscu JC (1995) The photosynthetic apparatus of phytoplankton from a perennially ice- covered Antarctic lake: acclimation to an extreme shade environment. Plant Cell Physiol 36: 253-263 Ouyang J, Shao X, Li J (2000) Indole‐3‐glycerol phosphate, a branchpoint of indole‐3‐acetic acid biosynthesis from the tryptophan biosynthetic pathway in Arabidopsis thaliana. Plant J 24: 327-334 Pfannschmidt T (2003) Chloroplast redox signals: how photosynthesis controls its own genes. Trends Plant Sci 8: 33-41 Pocock T, Vetterli A, Falk S (2011) Evidence for phenotypic plasticity in the Antarctic extremophile Chlamydomonas raudensis Ettl. UWO 241. J Exp Bot 62: 1169-1177 Possmayer M, Berardi G, Beall BFN, Trick CG, Hüner NPA, Maxwell DP (2011) Plasticity of the psychrophilic green alga Chlamydomonas raudensis (UWO 241) (Chlorophyta) to supraoptimal temperature stress. J Phycol 47: 1098-1109 Priscu JC, Wolf CF, Takacs CD, Fritsen CH, Laybourn-Parry J, Roberts JKM, Berry-Lyons W (1999) Carbon transformations in the water column of a perennially ice-covered Antarctic Lake. Biosci. 49: 997-1008 Raymond JA, Janech MG, Fritsen CH (2009) Novel ice-binding proteins from a psychrophilic Antarctic alga (Chlamydomonadaceae, Chlorophyceae). J. Phycol 45: 130-136

Sacksteder CA, Kramer DM (2000) Dark-interval relaxation kinetics (DIRK) of absorbance changes as a quantitative probe of steady-state electron transfer. Photosynth Res 66: 145- 158 Scheibe R (2004) Malate valves to balance cellular energy supply. 120: 21-26 Shimakawa G, Shaku K, Miyake C (2016) Oxidation of P700 in photosystem I is essential for the growth of cyanobacteria. Plant Physiol. 172: 1443-1450 Siaut M, Cuiné S, Cagnon C, Fessler B, Nguyen M, Carrier P, Beyly A, Beisson F, Triantaphylidès C, Li-Beisson Y (2011) Oil accumulation in the model green alga Chlamydomonas reinhardtii: characterization, variability between common laboratory strains and relationship with starch reserves. BMC Biotechnol 11: 7 Steinbeck J, Ross IL, Rothnagel R, Gäbelein P, Schulze S, Giles N, Ali R, Drysdale R, Sierecki E, Gambin Y (2018) Structure of a PSI–LHCI–cyt b6f supercomplex in Chlamydomonas reinhardtii promoting cyclic electron flow under anaerobic conditions. Proc Natl Acad Sci USA 115: 10517-10522 Strand DD, Livingston AK, Satoh-Cruz M, Froehlich JE, Maurino VG, Kramer DM (2015) Activation of cyclic electron flow by hydrogen peroxide in vivo. Proc Natl Acad Sci U S A 112: 5539-5544 Suorsa M (2015) Cyclic electron flow provides acclimatory plasticity for the photosynthetic machinery under various environmental conditions and developmental stages. Front Plant Sci 6: 800 Suzuki N, Bassil E, Hamilton JS, Inupakutika MA, Zandalinas SI, Tripathy D, Luo Y, Dion E, Fukui G, Kumazaki A (2016) ABA is required for plant acclimation to a combination of salt and heat stress. PloS one 11: e0147625 Szyska-Mroz B. , Cvetkovska M., Smith D.R., Possmayer M., Maxwell D.P. Hüner N.P.A. (2019) The Antarctic psychrophile, Chlamydomonas sp. UWO241, exhibits cold adapted protein kinases and thylakoid remodelling that confer a distinctive thylakoid protein phosphorylation pattern associated with a novel photosystem I / II energy distribution Plant Physiol. In Press Szyszka-Mroz B, Pittock P, Ivanov AG, Lajoie G, Hüner NP (2015) The Antarctic psychrophile, Chlamydomonas sp. UWO 241, preferentially phosphorylates a PSI-cytochrome b6/f supercomplex. Plant Physiol. 169: 717-736

Szyszka B, Ivanov AG, Huner NPA (2007) Psychrophily induces differential energy partitioning, photosystem stoichiometry and polypeptide phosphorylation in Chlamydomonas raudensis. Biochim Biophys Acta 1767: 789-800 Takabayashi A, Kishine M, Asada K, Endo T, Sato F (2005) Differential use of two cyclic electron flows around photosystem I for driving CO2-concentration mechanism in C4 photosynthesis. Proc Natl Acad Sci USA 102: 16898-16903 Takahashi H, Clowez S, Wollman F-A, Vallon O, Rappaport F (2013) Cyclic electron flow is redox-controlled but independent of state transition. Nature Comm 4: 1954 Tamoi M, Nagaoka M, Miyagawa Y, Shigeoka S (2006) Contribution of fructose-1, 6- bisphosphatase and sedoheptulose-1, 7-bisphosphatase to the photosynthetic rate and carbon flow in the Calvin cycle in transgenic plants. Plant Cell Physiol 47: 380-390 Terashima M, Petroutsos D, Hüdig M, Tolstygina I, Trompelt K, Gäbelein P, Fufezan C, Kudla J, Weinl S, Finazzi G (2012) Calcium-dependent regulation of cyclic photosynthetic electron transfer by a CAS, ANR1, and PGRL1 complex. Proc Natl Acad Sci USA 109: 17717- 17722 Thalmann M, Pazmino D, Seung D, Horrer D, Nigro A, Meier T, Kölling K, Pfeifhofer HW, Zeeman SC, Santelia D (2016) Regulation of leaf starch degradation by abscisic acid is important for osmotic stress tolerance in plants. Plant Cell 28: 1860-1878 Tohge T, Watanabe M, Hoefgen R, Fernie AR (2013) Shikimate and phenylalanine biosynthesis in the green lineage. Front Plant Sci 4: 62 Tzin V, Galili G (2010) The biosynthetic pathways for shikimate and aromatic amino acids in Arabidopsis thaliana. The Arabidopsis book/American Society of Plant Biologists 8 Valledor L, Weckwerth W (2014) An improved detergent-compatible gel-fractionation LC- LTQ-Orbitrap-MS workflow for plant and microbial proteomics. In Plant Proteomics. Springer, pp 347-358 van den Berg RA, Hoefsloot HC, Westerhuis JA, Smilde AK, van der Werf MJ (2006) Centering, scaling, and transformations: improving the biological information content of metabolomics data. BMC Gen 7: 142 Vanneste S, Friml J (2013) Calcium: the missing link in auxin action. Plants 2: 650-675 Wang X, Chang L, Wang B, Wang D, Li P, Wang L, Yi X, Huang Q, Peng M, Guo A (2013) Comparative proteomics of Thellungiella halophila leaves from plants subjected to salinity

reveals the importance of chloroplastic starch and soluble sugars in halophyte salt tolerance. Molec Cell Prot 12: 2174-2195 Wang X, Liu W, Xin C, Zheng Y, Cheng Y, Sun S, Li R, Zhu XG, Dai SY, Rentzepis PM, Yuan JS (2016) Enhanced limonene production in cyanobacteria reveals photosynthesis limitations. Proc Natl Acad Sci USA. 113: 14225-14230 Xu T, Venable J, Park SK, Cociorva D, Lu B, Liao L, Wohlschlegel J, Hewel J, Yates III J (2006) ProLuCID, a fast and sensitive tandem mass spectra-based protein identification program. In Molecular & cellular proteomics, Vol 5. Amer Soc Biochemistry Molecular Biology Inc, pp S174-S174 Yamori W, Makino A, Shikanai T (2016) A physiological role of cyclic electron transport around photosystem I in sustaining photosynthesis under fluctuating light in rice. Scientific Reports 6: 1-12 Zhang R, Cruz JA, Kramer DM, Magallanes-Lundback ME, Dellapenna D, Sharkey TD (2009) Moderate heat stress reduces the pH component of the transthylakoid proton motive force in light‐adapted, intact tobacco leaves. Plant Cell Environ. 32: 1538-1547

2.6 APPENDIX

Appendix Figure 2.1. Photosynthetic growth of UWO 241 under low and high salinity. Growth curves of UWO 241 under low (black) and high salinity (gray) (A). Growth was measured as optical density at 750 nm and maximum growth rates per day were calculated (µmax day-1, insert). Light response curves for net oxygen evolution rates (B). Oxygen evolution was measured in mid-log phase cultures. No significant difference was observed between HS and LS cultures (two tailed t-test, unequal variance; n=3).

Appendix Figure 2.2. Photosynthetic properties of UWO 241 under various light intensities. The PSII operating efficiency (A), NPQ (B) and LEF (C) of C. sp. UWO 241 grown in low-salt (black, squares) and high-salt (grey, circles) were measured by saturation-pulse chlorophyll

fluorescence yield changes. The τECS (D) of C. sp. UWO 241 were estimated from dark interval relaxation kinetics (DIRK) of electrochromic shift (ECS). *Values were statistically significant between HS and LS cultures (p<0.005; two tailed t-test, unequal variance; n=3).

Y(PSII) Y(NPQ) Y(NO) Y(PSI) (Y(NA) Y(ND) A B 1.00 1.00

0.75 0.75

0.50 0.50

0.25 0.25

PSII Energy (%) Partitioning PSI Energy (%) Partitioning 0.00 0.00 Control HS Control HS

Appendix Figure 2.3. PSI and PSII energy partitioning in cultures of UWO 241 under low salt (LS) versus high salinity (HS) conditions. A. PSII energy partitioning. B. PSI energy partitioning. Y(PSII), Photochemical yield of PSII; Y(NO), non-regulated energy dissipation; Y(NPQ), non-photochemical quenching; Y(PSI), Photochemical yield of PSI; Y(ND), donor-side limitation of PSI; Y(NA), acceptor-side limitation of PSI.

Appendix Figure 2.4. Maximum likelihood tree of all LHCA protein homologues identified in the UWO 241 transcriptome from HS-grown cultures. Bolded sequences represent cDNA contigs recovered from the UWO 241 transcriptome. Asterix indicates LHCA protein identified in proteome of the UWO 241 PSI supercomplex. Numbers are bootstrap values from 500 replicate trees. CR- Chlamydomonas reinhardtii, ARA- Arabidopsis thaliana

Miscellaneous Up-regulated Cell cycle and structure Down-regulated Repair Oxidative Stress Protein Synthesis Other Metabolism Carbohydrate metabolism and storage Photosynthesis Membrane & Transport

0 5 10 15 20 number of proteins

Appendix Figure 2.5. Functional categories of proteins up-regulated (Black) and down-regulated (Gray) under high salinity in Chlamydomonas sp. UWO 241.

ATP content 1.0 *

0.8 cells 6 0.6

0.4

0.2 ATP (nmol)/10

0.0

UWO241-LSUWO241-HS

Appendix Figure 2.6. Cellular ATP content in UWO 241 grown under low salinity (UWO 241- LS) versus high salinity (UWO 241-HS) growth conditions. (n=3; *, statistical difference, p<0.05).

Appendix Figure 2.7: Heat map showing the relative changes in primary metabolite abundances in UWO 241 cultures grown under high or low salt conditions. The analysis includes all quantified metabolites. For each growth condition, three biological replicates are represented using a color-based metabolite profile (log2 normalized metabolite abundance; red – high abundance; blue – low abundance). Hierarchical clustering is based on Euclidean distances and Ward’s linkage.

50 A - C. reinhardtii

30 A820 (mV) A820

B- UWO241 LS

40 A820 (mV) A820

C UWO241 HS

A820 (mV) A820 40

0 0 2 4 6 8 10 12 14 time (s)

Appendix Figure 2.8. Representative traces of P700 photooxidation kinetics. Traces represent mean±S.D. of three technical replicates

50 UWO241-LS UWO241-HS 40

cells) 30 6

20 g per 10 Starch content

µ 10 (

UWO241-LSUWO241-HS Appendix Figure 2.9: Starch content of UWO 241 under low salt (black) and high salt (gray) conditions. Log-phase cultures were used, and starch was quantified using commercial starch kit utilizing amyloglucosidase enzyme. (n=3).

Appendix Table 2.1. Composition of PSI and LHCI subunits in UWO 241. Subunits of PSI and LHCI identified in the proteome of band 3 and band 4 of thylakoid sucrose density gradient, and whole cells under low (LS) and high (HS) salinity are shown.

Protein Band 3 Band 4 Whole cell Whole Cell

LS HS

PSI PsaA X X PsaB X X X PsaC PsaD X X X X PsaE X X X X PsaF X X X X PsaH X X X X PsaK X X X X PsaL X X X PsaN X X X LHCI Lhca1 Lhca2 Lhca3 X X X X Lhca4 Lhca5 X X X X Lhca6 Lhca7 Lhca8 Lhca9

Appendix Table 2.2. Down-regulated proteins in UWO 241-HS. Identified proteins are divided into functional categories and organelle localization (Sub. Loc.) is provided based on Uniprot prediction. Number of proteins belonging to each category is provided in parenthesis. (FC- fold change)

PROTEIN NAME Uniprot SUB. LOC. PROTEIN FUNCTION FC p-value KB MEMBRANE AND TRANSPORT(5) ABC transporter C family member 3 Q54JR2 Transmembrane movement 1.56 0.0031 Dicarboxylate/tricarboxylate Q9C5M0 Mitochondria Transport of 84ecarboxylase 1.40 0.0285 transporterClathrin heavy chain 1 Q2RBN7 Coated vesicular transport 2.19 0.00003 Clathrin heavy chain 2 Q2QYW2 Coated vesicular transport 2.65 0.0039 V-type proton ATPase catalytic Q38676 Vacuole Acidification of endomembrane 2.39 0.0001 subunit A isoform 1 by proton transport PHOTOSYNTHESIS (7)

NAD(P)H-quinone oxidoreductase Q9T0A4 Chloroplast Electron transport to quinone 2.37 0.0024 subunit S Photosystem II D2 protein A6YG76 Chloroplast PSII reaction center protein 2.04 0.0169

Photosystem I P700 chlorophyll a P09144 Chloroplast PSI reaction center protein 1.88 0.0020 apoprotein A2 Photosystem I reaction center subunit Q9SBN5 Chloroplast PSI antenna binding 1.85 0.0205 N Oxygen-evolving enhancer protein 1 Q9SBN6 Chloroplast Stabilization of OEC 1.41 0.0102

Oxygen-evolving enhancer protein 3 P12852 Chloroplast Stabilization of OEC, PSII 1.29 0.0033 biogenesis Geranylgeranyl diphosphate Q9CA67 Chloroplast Tocopherol and Chlorophyll reductase synthesis CARBOHYDRATE METABOLISM AND STORAGE (4) Aldehyde-alcohol dehydrogenase P0A9Q7 Ethanol Biosynthesis 5.33 0.0040

Pyruvate Carboxylase O93918 Cytoplasm Gluconeogenesis 2.34 0.0134

Malate Dehydrogenase P46489 Chloroplast Malate Shunt 2.13 0.0060

Biotin Carboxylase O04983 Chloroplast Lipid Biosynthesis 1.43 0.0168

OTHER METABOLISMS (5)

24-methylenesterol C- Q39227 Chloroplast Steroid Biosynthesis 1.33 0.0035 methyltransferase 2 Phosphoserine aminotransferase 1 Q96255 Chloroplast Serine/ Amino acid Biosynthesis 2.16 0.011

Carbamoyl-phosphate synthase large B9EXM2 Chloroplast Arginine/ Amino acid 1.63 0.0057 chain Biosynthesis Argininosuccinate lyase P22675 Arginine/ Amino acid 1.62 0.034 Biosynthesis NADH dehydrogenase [ubiquinone] 1 Q9FIJ2 Mitochondria Respiration 1.65 0.012 alpha subcomplex subunit 2 PROTEIN SYNTHESIS (4) Ribosomal protein L9 Q7UZJ1 Chloroplast Translation 2.83 0.0028

Ribosomal protein L29 Q46IR7 Translation 1.92 0.0116

Translocon-associated protein subunit Q5E9E4 ER Calcium binding chaperone of 1.85 0.0277 beta translocated proteins Protein disulfide-isomerase like 2-2 Q9MAU6 ER Protein folding catalyst 1.48 0.0216

OXIDATIVE STRESS (1) Superoxide dismutase [Mn] Q42684 Mitochondria ROS detoxification 2.35 0.0134

REPAIR (3) ATP-dependent A3DH20 Double stranded DNA break 2.02 0.0190 helicase/deoxyribonuclease subunit B repair HSP70-HSP90 organizing protein 3 Q9STH1 Nucleus Heat Acclimation/chaperone 1.61 0.0136 association Ribonuclease 3 Q1D5X9 Nucleus Nuclease activity on double 1.46 0.0024 stranded RNA CELL CYCLE AND STRUCTURE (5) Dynamin-related protein 1C Q8LF21 Cytoskeleton Microtubule binding, cell 1.73 0.0209 division Dynamin-related protein 1A P42697 Cytoskeleton Cell wall maintenance 2.25 0.0291

Dynamin-2B Q9LQ55 Cytoskeleton Clathrin dependent endocytosis 1.49 0.0178

Tubulin alpha-2 chain P09205 Cytoskeleton Microtubule formation 2.48 0.0054

Tubulin alpha-1A chain P50258 Cytoskeleton Microtubule formation 1.58 0.0075

MISCELLANEOUS (4) Cytosol non-specific dipeptidase P15288 Cytoplasm Broad dipeptidase activity 2.49 0.0046

Calcium/calmodulin-dependent A8WXF6 Calcium dependent Ser/Thr 2.10 0.0205 protein kinase type II kinase activity Microbial collagenase Q8D4Y9 Peptidase 1.87 0.0294

Putative F-box/kelch-repeat protein Q9SY96 Ubiquitination of proteins 1.774 0.0088

Appendix Table 2.3. Up-regulated proteins in UWO 241-HS. Identified proteins are divided into functional categories and organelle localization (Sub. Loc.) is provided based on Uniprot prediction. Number of proteins belonging to each category is provided in parenthesis. (FC- fold change)

PROTEIN NAME Uniprot SUB. LOC. PROTEIN FUNCTION FC p-value KB MEMBRANE AND TRANSPORT (3) Probable cation-transporting ATPaseF P9WPS9 Cytoplasm Sodium ion pump 3.95 0.0011

Import receptor subunit TOM40-1 Q9LHE5 Mitochondria Pore formation 1.8 0.01 ABC transporter I family member 6 Q9CAF5 Chloroplast Transmembrane movement of 1.56 0.003 substance, chloroplast PHOTOSYNTHESIS (12) differentiation ATP synthase epsilon P07891 Chloroplast ATP synthesis 3.8 0.0218

ATP-dependent zinc metalloprotease 1 Q5Z974 Chloroplast PSII repair, CEF 2.1 0.0004

ATP-dependent zinc metalloprotease 2 O80860 Chloroplast PSII repair, thylakoid 1.69 0.009 organization, CEF Photosystem II stability/assembly factor O82660 Chloroplast PSII and cytb559 biogenesis 1.45 0.0042 HCF136 RubisCo large subunit binding protein Q42694 Chloroplast Assembly of RubisCO 1.87 0.0005 alpha RubisCo large subunit binding protein P21241 Chloroplast Assembly of RubisCO 1.77 0.0162 beta Transketolase Q43848 Chloroplast Calvin cycle 1.38 0.023

Ribulose bisphosphate carboxylase large P08211 Chloroplast Calvin cycle 1.35 0.0069 chain Ribose-5-phosphate isomerase 3 Q9S726 Chloroplast Calvin cycle 2.29 0.0053

Phosphoribulokinase P19824 Chloroplast Calvin cycle 1.26 0.0005

Fructose-1,6-bisphosphatase P9WN21 Chloroplast Calvin cycle 5.08 0.0006

15-cis-phytoene desaturase Q07356 Chloroplast Carotenoid biosynthesis 1.31 0.0079

CARBOHYDRATE METABOLISM AND STORAGE (7) Aconitate Hydratase Q99KI0 Mitochondria TCA cycle 1.54 0.0308

Fructose bis-phosphate aldolase O52402 Cytoplasm Gluconeogenesis 1.48 0.0253

Glucose-1-phosphate P55233 Chloroplast Starch synthesis 1.55 0.0031 adenylyltransferase large subunit Starch Synthase 1 Q43784 Chloroplast Starch synthesis 1.47 0.0045

Glycerol-3-phosphate dehydrogenase P52425 Cytosol Glycerol biosynthesis 6.28 0.00001 [NAD+] Glycerol-3-phosphate dehydrogenase Q949Q0 Chloroplast TAG biosynthesis 1.7 0.0094 [NAD+] Inorganic soluble pyrophosphatase 1 Q93Y52 Chloroplast Recycling inorganic 1.8 0.0063 pyrophosphate OTHER METABOLISMS (13) DAHP Synthase Q00218 Chloroplast Shikimate Pathway 2.28 0.0285

Phospho-2-dehydro-3-deoxyheptonate P29976 Chloroplast Shikimate Pathway 1.5 0.02181 aldolase1 Chorismate Synthase 1 Q42884 Chloroplast Shikimate Pathway 1.93 0.0097

Indole-3-glycerol phosphate synthase P49572 Chloroplast Tryptophan Biosynthesis 2.64 0.0256

Phosphomethylpyrimidine synthase O82392 Chloroplast Thiamine diphosphate 2.15 0.002 biosynthesis, Cofactor biosynthesis GDP-mannose 3,5-epimerase 2 Q2R1V8 Cytoplasm Vitamin C biosynthesis 1.37 0.0005

Glycine dehydrogenase O49954 Mitochondria Glycine cleavage system, 1.37 0.0061 (decarboxylating) Photorespiration Aspartate-semialdehyde dehydrogenase Q55512 Cytoplasm Amino acid biosynthesis 1.81 0.023

Glutamate--glyoxylate aminotransferase Q9S7E9 Peroxisome Amino acid degradation 1.80 0.0027 2 S-adenosylmethionine synthase Q4JIJ3 Cytoplasm Amino acid biosynthesis 1.58 0.00001

Ketol-acid reductoisomerase Q05758 Chloroplast Amino acid biosynthesis 1.42 0.0236

Methionine synthase A8HYU5 Cytoplasm Amino acid biosynthesis 1.31 0.0009

Sulfate adenylyltransferase Q0V6P9 Cytoplasm Amino acid biosynthesis 1.93 0.0098

PROTEIN SYNTHESIS (15) Ribosomal protein L19 B8HT34 Chloroplast Translation 3.09 0.0051

Ribosomal Protein L22 Q84U21 Chloroplast Translation 1.87 0.0023

Ribosomal Protein L23 Q8HTL3 Chloroplast Translation 1.56 0.0011

Ribosomal protein L13 O48513 Chloroplast Translation 1.76 0.0327

40S ribosomal protein S23-2 P49201 Cytoplasm Translation 2.69 0.0133

40S ribosomal protein S14 P46295 Cytoplasm Translation 2.31 0.0027

Eukaryotic translation initiation factor 2 Q9SIZ2 Cytoplasm/ Translation 1.74 0.0065 subunit alpha homolog Nucleus Elongation factor 3 O94489 Cytoplasm Translation 1.73 0.0004

Elongation factor Tu P17746 Chloroplast Translation 1.64 0.0076

Ribosomal protein S26 P62856 Cytoplasm Translation 1.60 0.0126

Ribosomal Protein L3 P35684 Cytoplasm Translation 1.59 0.0231

Ribosomal Protein L30-2 Q8VZ19 Cytoplasm Translation 1.51 0.0149

Eukaryotic initiation factor 4A-11 Q40465 Cytoplasm Translation 1.59 0.0002

Eukaryotic translation initiation factor Q9AXQ5 Translation 1.49 0.0145 5A-2 60S acidic ribosomal protein P0 P50346 Translation 1.38 0.0010

OXIDATIVE STRESS (3) Putative heme-binding peroxidase A4R606 ROS detoxification 5.24 0.0015

Superoxide dismutase [Fe] P22302 Chloroplast ROS detoxification 3.15 0.0027

Obg-like ATPase 1 Q9SA73 Cytoplasm Lipid peroxidation 2.10 0.0122

REPAIR (1) DNA mismatch repair protein MutS Q7V9M5 Mismatch Repair 2.72 0.00261

CELL CYCLE AND STRUCTURE (1) Cell division cycle protein 48 homolog Q96372 Cell Division 1.68 0.0045

Appendix Table 2.4. All positively identified primary metabolites in UWO 241. Identification of metabolites is based on GC-MS spectra and retention time. Metabolites which are present in significantly higher amounts in high salt grown cultures are highlighted in green (FC>2; FDR- adjusted p<0.05). Metabolites which are present in significantly lower amounts in high salt grown cultures are highlighted in red (FC>2; FDR-adjusted p<0.05).

Metabolite KEGG Chemical Class log2(FC) p-value sulfuric acid C00059 Acid 1.9036 0.033685 cysteine C00097 Amino acid -1.5236 1.21E-05 beta-alanine C00099 Amino acid -0.7103 0.002515 alanine-alanine C00993 Amino acid -1.2634 0.002702 alanine C00041 Amino acid -1.422 0.008152 asparagine C00152 Amino acid -1.5839 0.011711 methionine sulfoxide C02989 Amino acid -1.643 0.015325 isoleucine C00407 Amino acid -0.89692 0.018012 cystine C01420 Amino acid -0.93734 0.018638 citrulline C00327 Amino acid -2.2366 0.023489 histidine C00135 Amino acid -4.6082 0.042118 glutamine C00064 Amino acid -1.5487 0.065586 glycine C00037 Amino acid -0.73068 0.073107 5-hydroxynorvaline, pentahomoserine Amino acid -0.65158 0.075262

4-aminobutyric acid C00334 Amino acid -0.38668 0.27338 3-aminoisobutyric acid C05145 Amino acid -0.3345 0.38567 aspartic acid C00049 Amino acid -0.19633 0.57033 cysteine-glycine C01419 Amino acid -0.62178 0.61831 proline C00148 Amino acid 4.7609 4.68E-05 phenylalanine C00079 Amino acid -1.7007 0.003189 glutamic acid C00025 Amino acid -1.7161 0.003219 tyrosine C00082 Amino acid -2.2116 0.005018 ornithine C00077 Amino acid -3.8106 0.005184 lysine C00047 Amino acid -4.8771 0.009432 valine C00183 Amino acid -1.2949 0.01344 tryptophan C00078 Amino acid -1.4968 0.017354 leucine C00123 Amino acid -0.87848 0.04099 methionine C00073 Amino acid -1.1444 0.20888 threonine C00188 Amino acid -0.71743 0.20907

serine C00065 Amino acid -0.58687 0.26196 oxoproline, pyroglutamic acid C01879 Amino acid -0.27963 0.38632 norvaline C01826 Amino acid -0.37303 0.44916 O-acetylserine C00979 Amino acid -0.53416 0.58989 N-acetylglutamate C00624 Amino acid, byproduct -1.4239 0.004714 glycyl-proline Amino acid, byproduct -0.85676 0.20112

N-acetylornithine C00437 Amino acid, byproduct -0.641 0.26131 N-acetylaspartic acid C01042 Amino acid, byproduct -0.26014 0.61357 glycyl tyrosine Amino acid, byproduct -0.49218 0.69894

n-epsilon-trimethyllysine C03793 Amino acid, byproduct -0.27774 0.73808 2,4-diaminobutyric acid C03283 Amino acid, derivative -1.497 0.001906 guanidinosuccinate C03139 Amino acids, derivative 2.7405 0.000409 2-aminobutyric acid, N-methyl-alanine C02721 Amino acids, derivative 0.65503 0.089108 2-ketoglucose dimethylacetal, D-glucosone Carbohydrate -2.3512 0.00255 xylonolactone C02266 Carbohydrate 0.30723 0.60649 trehalose C01083 Carbohydrate, disaccharide 0.15995 0.69592 ketohexose (Tagatose) C00795 Carbohydrate, monosaccharide -0.28621 0.26141 N-acetyl-D-galactosamine C05021 Carbohydrate, monosaccharide 0.68096 0.36352 sucrose C00089 Carbohydrate, monosaccharide 4.1851 0.012685 xylose C00181 Carbohydrate, monosaccharide -1.4883 0.034587 fructose C02336 Carbohydrate, monosaccharide -1.0881 5.79E-02 glucose C00221 Carbohydrate, monosaccharide 0.84823 0.19743 n-acetyl-d-hexosamine C03878 Carbohydrate, monosaccharide -0.76489 0.36213 ribose C00121 Carbohydrate, monosaccharide 0.17182 0.82713 erythrose C01796 Carbohydrate, monosaccharide 0.021311 0.93894 raffinose C00492 Carbohydrate, trisaccharide 0.56229 0.086431 maltotriose C01835 Carbohydrate, trisaccharide 0.20192 0.47094 melezitose C08243 Carbohydrate, trisaccharide 0.23215 0.74992 lactose C00243 Carbohydrates, disaccharides 1.3114 0.095967 N-acetylmannosamine C00645 Carbohydrates, monosaccharide -0.93289 0.25098 erythronic acid lactone C02341 Carboxylic acid ester -1.4731 0.013289 1-monopalmitin C01885 Lipid, glycerol derivative 1.0142 0.053251 1-monoolein Lipid, glycerol derivative 0.88862 0.10514

1-monostearin D01947 Lipid, glycerol derivative -0.08456 0.60961 ergosterol C01694 Lipid, sterol 4.0604 0.055931 myristic acid, tetradecanoic acid C06424 Lipids, fatty acid, saturated 0.83412 0.011937 pentadecanoic acid C16537 Lipids, fatty acid, saturated -0.72024 0.043494

palmitic acid C00249 Lipids, fatty acid, saturated 0.27594 0.17162 stearic acid C01530 Lipids, fatty acid, saturated -0.24951 0.18977 pelargonic acid, nonanoic acid C01601 Lipids, fatty acid, saturated -0.27423 0.37938 lauric acid, dodecanoic acid C02679 Lipids, fatty acid, saturated 0.23044 0.62134 oleic acid C00712 Lipids, fatty acid, unsaturated -0.19992 0.58631 palmitoleic acid, hexadecanoic acid C08362 Lipids, fatty acid, unsaturated -0.46713 0.89761 heptadecanoic acid, margaric acid Lipids, fatty acids, saturated 1.546 0.09097

cerotinic acid Lipids, fatty acids, saturated 0.2003 0.32045

capric acid, decanoic acid C01571 Lipids, Fatty acids, saturated -0.31306 0.43639 cis-gondoic acid, isosenoic acid C16526 Lipids, Fatty acids, unsaturated 0.92074 0.007523 linoleic acid C01595 Lipids, Fatty acids, unsaturated 0.94152 0.008666 linolenic acid C06427 Lipids, Fatty acids, unsaturated -0.76121 0.085481 phytol C01389 Lipids, isoprenoids 3.6521 0.018108 stigmasterol C05442 Lipids, sterol -1.3717 0.32312 hydroxycarbamate N-containing -0.01 0.84584

uric acid C00366 N-containing -2.3549 0.012194 2-hydroxypyrazinyl-2-propenoic acid ethyl ester N-containing -0.19844 0.52948

urea C00086 N-containing -0.14154 0.85024 2-picolinic acid C10164 N-containing, alkaloid -2.0521 0.32111 lactamide N-containing, amide -0.17792 0.375

butyrolactam N-containing, amide -1.0986 4.62E-05 epsilon-caprolactam C06593 N-containing, amide -0.52274 0.37827 cyclohexylamine C00571 N-containing, amine -0.34572 0.30004 putrescine C00138 N-containing, amines -3.5847 0.001276 maleimide C07272 N-containing, imide -0.59643 0.34092 guanine C00242 N-containing -1.009 0.17819 adenosine C00212 N-containing, -1.9445 0.099988 inosine 5'-monophosphate, inosinic acid C00130 N-containing 1.9176 0.21012 2'-deoxyguanosine C00330 N-containing 0.31645 0.24821 5'-deoxy-5'-methylthioadenosine C00170 N-containing -0.29362 5.15E-01 inosine C00294 N-containing -0.15072 0.8453 xanthosine C01762 N-containing -0.11131 0.39975 pseudo-uridine C02067 N-containing -1.7419 0.011404 uridine C00299 N-containing -0.49148 0.34373 5-methyluridine, ribothymidine N-containing 0.036994 0.80971

UDP-N-acetylglucosamine C00043 N-containing -0.731 0.004359 adenosine-5-monophosphate C00020 N-containing, P-containing -1.838 0.003937

3'-adenylic acid, Adenosine 3-monophosphate C01367 N-containing, P-containing 0.86286 0.15054 adenine C00147 N-containing -1.7093 0.00287 cytidine-5-monophosphate C00055 N-containing -2.2385 0.00375 cytosin C00380 N-containing -0.35963 0.49354 hypoxanthine C00262 N-containing, Purine derivative -1.4848 0.044797 uracil C00106 N-containing, Pyrimidine nucleobase -1.2572 0.03763 thymine C00178 N-containing, Pyrimidine nucleobase -0.6117 0.1543 thymidine C00214 N-containing, Pyrimidine nucleoside -0.45464 0.35538 hydroxylamine C00192 N-containing -0.45436 0.22993 xanthine C00385 N-containing -0.75414 0.10379 parabanic acid C00802 Organic acid -1.3733 0.000303 isothreonic acid C00639 Organic acid -2.6604 0.001234 2-ketoisocaproic acid C00233 Organic acid -2.6763 0.004356 phenylpyruvate, phenylpyruvic acid C00166 Organic acid -1.3252 0.005513 4-hydroxybutyric acid C00989 Organic acid -0.89867 0.007894 2-hydroxyhexanoic acid Organic acid -1.1636 0.015951 pyrrole-2-carboxylic acid C05942 Organic acid 1.7601 0.019385 2-hydroxyglutaric acid C02630 Organic acid -1.8571 0.032588 orotic acid C00295 Organic acid -0.63526 0.13219 phosphogluconic acid C00345 Organic acid -0.76785 0.45608 erythronic acid Organic acid, Carboxylic acid 2.5583 0.001159 phosphoenolpyruvate C00074 Organic acid, Carboxylic acid -2.0419 0.001212 3-phosphoglycerate C00597 Organic acid, carboxylic acid -1.5661 2.26E-03 malic acid, malate C00711 Organic acid, Carboxylic acid -2.9173 0.004016 fumaric acid, fumarate C00122 Organic acid, Carboxylic acid -2.8632 0.0043 alpha-ketoglutarate C00026 Organic acid, Carboxylic acid -3.1097 0.004574 gluconic acid C00800 Organic acid, Carboxylic acid -1.0445 0.007016 pyruvic acid C00022 Organic acid, Carboxylic acid -3.117 0.007106 glycolic acid C00160 Organic acid, Carboxylic acid -0.81919 0.007484 succinic acid C00042 Organic acid, Carboxylic acid -1.7759 0.011196 oxalic acid C00209 Organic acid, Carboxylic acid -0.97491 0.023528 citramalic acid C00815 Organic acid, Carboxylic acid -1.7249 0.030439 alpha-aminoadipic acid C00956 Organic acid, Carboxylic acid -0.53697 0.17605 aminomalonate C00872 Organic acid, Carboxylic acid -0.72502 0.19176 lactic acid C00186 Organic acid, Carboxylic acid -0.43693 0.20046 benzoic acid C00180 Organic acid, Carboxylic acid -0.11954 0.39334 citric acid C00158 Organic acid, Carboxylic acid 0.18216 0.55282

3-hydroxy-3-methylglutaric acid C03761 Organic acid, carboxylic acid -0.24678 0.80945 ethanol phosphate P-containing -2.3425 0.18557

pyrophosphate (diphosphate) C00013 P-containing 0.077274 0.78319 propane-1,3-diol C02457 Polyol -0.02244 0.93984 glycerol C00116 Polyol 3.1322 0.000117 glycerol 2-phosphate C02979 Polyol -1.3775 0.004539 glycerol-3-galactoside C05401 Polyol -1.5253 0.005353 glycerol-1-phosphate C03189 Polyol -0.94915 0.030117 glyceric acid C00258 Polyol 0.29534 0.21221 threonic acid C01620 Sugar acid -2.3149 0.001672 galactinol C01235 Sugar alcohol -3.0243 0.000464 pentitol, xylitol D00061 Sugar alcohol 1.3426 0.001886 6-deoxyglucitol, sorbitol C00794 Sugar alcohol -2.8559 0.010357 mannitol C00392 Sugar alcohol -1.7028 0.09623 myo-inositol C00137 Sugar alcohol -0.35546 0.1282 isothreitol C16884 Sugar alcohol 0.76905 0.55649 monomyristin, Myristoyl glycerol Sugar alcohol -0.16368 0.64949

lactitol C13542 Sugar alcohol -0.24374 0.66554 lyxitol (arabinol) C00532 Sugar alcohol 0.005254 0.87387 ribose-5-phosphate C00117 Sugar phosphate -1.5405 0.013279 fructose-1-phosphate C01094 Sugar phosphate -2.0115 1.76E-02 inositol-4-monophosphate C03546 Sugar phosphate 0.79069 0.031381 fructose-1,6-bisphosphate C05378 Sugar phosphate -2.7527 0.035676 galactose-6-phosphate C01113 Sugar phosphate 1.3478 0.057026 ribulose-5-phosphate C00199 Sugar phosphate -1.0437 0.082425 glucose-1-phosphate C00103 Sugar phosphate 1.0028 0.15246 hexose-6-phosphate C02965 Sugar phosphate -0.67702 0.23873 fructose-6-phosphate C05345 Sugar phosphate -0.65442 0.27619 glucose-6-phosphate C00092 Sugar phosphate -0.54222 0.31425 mannose-6-phosphate C00275 Sugar phosphate 0.076688 0.60776 ascorbic acid C00072 Vitamins, cofactors -0.69859 0.031888 dehydroascorbic acid C05422 Vitamins, cofactors -2.4945 0.000199 nicotinamide C00153 Vitamins, cofactors -1.4902 0.002861 pyridoxal-5-phosphate C00018 Vitamins, cofactors -1.0647 0.011951 pantothenic acid C12276 Vitamins, cofactors -1.0914 0.16118 nicotinic acid C00253 Vitamins, cofactors 0.039776 0.68392 tocopherol alpha- C00376 Vitamins, cofactors 3.2516 0.060112

95 tocopherol gamma- C02483 Vitamins, cofactors 0.27428 0.39509 tocopherol beta C14152 Vitamins, cofactors 0.011064 0.93487

Appendix Table 2.5. Metabolites present in higher abundance in UWO 241-HS. Only metabolites which were positively identified based on their GC-MS spectra and retention times were taken into consideration. In all cases, metabolite abundance is significantly different between low and high salt grown algal cultures (t-test; FDR-adjusted p<0.05) and the fold change (FC) is >2. Identified metabolites are divided into chemical categories and pathways based on KEGG Pathway analysis.

Chemical Metabolite KEGG Pathway FC p-value category

Glycerol C00116 Polyol Glycerolipid metabolism 8.77 0.00012

Sucrose C00089 Carbohydrate Sugar and starch metabolism 18.19 0.01269

Proline C00148 Amino acid Amino acid metabolism 27.11 0.00005

Guanidinosuccinate C03139 Organic acid Amino acid metabolism 6.68 0.00041

Pyrrole-2-carboxylic acid C05942 Organic acid Amino acid metabolism 3.39 0.01939

Sulfur metabolism, Amino acid Sulfuric acid C00059 Acid 3.74 0.03369 metabolism

Erythronic acid C21593 Organic acid Ascorbic acid degradation 5.89 0.00116

Lipids, Phytol C01389 Chlorophyll a degradation product 12.57 0.01811 isoprenoid

Xylitol D00061 Sugar alcohol Xylose degradation 2.54 0.00189

Appendix Table 2.6. Metabolites present in lower abundance in UWO 241-HS. Only metabolites which were positively identified based on their GC-MS spectra and retention times were taken into consideration. In all cases, metabolite abundance is significantly different between low and high salt grown algal cultures (t-test; FDR-adjusted p<0.05) and the fold change (FC) is >2. Identified metabolites are divided into chemical categories and pathways based on KEGG Pathway analysis.

Metabolite KEGG Chemical category Pathway FC p-value Cysteine C01420 Amino acid Amino acid metabolism 5.75 0.00001 Alanine-alanine C00993 Amino acid Amino acid metabolism 2.40 0.00270 Amino acid metabolism, Carbon Alanine C00041 Amino acid 2.68 0.00815 fixation Asparagine C00152 Amino acid Amino acid metabolism 3.00 0.01171 Methionine sulfoxide C02989 Amino acid Amino acid metabolism 3.12 0.01533 Citrulline C00327 Amino acid Amino acid metabolism 4.71 0.02349 Histidine C00135 Amino acid Amino acid metabolism 24.39 0.04212 Phenylalanine C00079 Amino acid Amino acid metabolism 3.25 0.00319 Amino acid metabolism, Porphyrin Glutamic acid C00025 Amino acid and chlorophyll metabolism 3.29 0.00322 Nitrogen metabolism Tyrosine C00082 Amino acid Amino acid metabolism 4.63 0.00502 Ornithine C00077 Amino acid Amino acid metabolism 14.03 0.00518 Putrescine C00138 N-containing Amino acid metabolism 12.00 0.00128 Lysine C00047 Amino acid Amino acid metabolism 29.39 0.00943 Valine C00183 Amino acid Amino acid metabolism 2.45 0.01344 Tryptophan C00078 Amino acid Amino acid metabolism 2.82 0.01735 Amino acid N-acetylglutamate C00624 Amino acid metabolism 2.68 0.00471 (byproduct) Amino acid 2,4-diaminobutyric acid C03283 Amino acid metabolism 2.82 0.00191 (derivative) D-glucosone Carbohydrate 5.10 0.00255 Amino sugar and nucleotide sugar Xylose C00181 Carbohydrate 2.81 0.03459 metabolism Erythronic acid lactone C02341 Carboxylic acid ester 2.78 0.01329 Nicotinate and nicotinamide Nicotinamide C00153 Cofactors 2.81 0.00286 metabolism Butyrolactam N-containing Caprolactam degradation 2.14 0.00005 Adenine C00147 N-containing Purine metabolism 3.27 0.00287 Cytidine-5-monophosphate C00055 N-containing Pyrimidine metabolism 4.72 0.00375

Pseudo-uridine C02067 N-containing Pyrimidine metabolism 3.34 0.01140 Uric acid C00366 N-containing Purine metabolism 5.12 0.01219 Uracil C00106 N-containing Pyrimidine metabolism 2.39 0.03763 Hypoxanthine C00262 N-containing Purine metabolism 2.80 0.04480 N-containing, P- Adenosine-5-monophosphate C00020 Purine metabolism 3.58 0.00394 containing Parabanic acid C00802 Organic acid Purine metabolism 2.59 0.00030 Glycolysis / Gluconeogenesis, TCA cycle, Amino acid metabolism, Phosphoenolpyruvate C00074 Organic acid 4.12 0.00121 Pyruvate metabolism, Carbon fixation Isothreonic acid C00639 Organic acid Ascorbate metabolism 6.32 0.00123 Threonic acid C01620 Organic acid Ascorbate metabolism 4.98 0.00167 Carbon, fixation, Glycolysis / Gluconeogenesis, Pentose phosphate 3-phosphoglycerate C00597 Organic acid 2.96 0.00226 pathway, Amino acid metabolism, Glycerolipid metabolism TCA cycle, pyruvate metabolism, Malic acid C00711 Organic acid 7.55 0.00402 carbon fixation TCA cycle, Oxidative phosphorylation, Pyruvate Fumaric acid C00122 Organic acid metabolism, Amino acid 7.28 0.00430 metabolism, Nicotinate and nicotinamide metabolism Valine, leucine and isoleucine 2-ketoisocaproic acid C00233 Organic acid biosynthesis 6.39 0.00436 Glucosinolate biosynthesis TCA cycle, Ascorbate metabolism alpha-ketoglutarate C00026 Organic acid 8.63 0.00457 Amino acid metabolism Phenylpyruvate C00166 Organic acid Amino acid metabolism 2.51 0.00551 Gluconic acid C00800 Organic acid Ascorbate metabolism 2.06 0.00702 Glycolysis / Gluconeogenesis, TCA cycle, Pentose phosphate pathway, Ascorbate metabolism, Amino acid Pyruvic acid C00022 Organic acid 8.68 0.00711 metabolism, Carbon fixation, Nicotinate and nicotinamide metabolism TCA cycle, Oxidative phosphorylation, Amino acid Succinic acid C00042 Organic acid 3.42 0.01120 metabolism, Pyruvate metabolism, Sulfur metabolism

2-hydroxyhexanoic acid Organic acid 2.24 0.01595 Glycolysis / Gluconeogenesis, TCA Citramalic acid C00815 Organic acid 3.31 0.03044 cycle Glycerol 2-phosphate C02979 Polyol Glycerolipid metabolism 2.60 0.00454 Glycerol-3-galactoside C05401 Polyol Glycerolipid metabolism 2.88 0.00535 Phosphoethanolamine C00346 N-containing Glycerolipid metabolism 10.13 0.00004 Galactinol C01235 Sugar alcohol Galactose metabolism 8.14 0.00046 Fructose and mannose metabolism, 6-deoxyglucitol sorbitol C00794 Sugar alcohol 7.24 0.01036 Galactose metabolism Carbon fixation, Pentose phosphate Ribose-5-phosphate C00117 Sugar phosphate 2.91 0.01328 pathway, Purine metabolism Amino acid metabolism, Fructose Fructose-1-phosphate C01094 Sugar phosphate 4.03 0.01757 and mannose metabolism Glycolysis / Gluconeogenesis, Fructose-1,6-bisphosphate C05378 Sugar phosphate Pentose phosphate pathway, 6.74 0.03568 Fructose and mannose metabolism Pyridoxal-5-phosphate C00018 Vitamin, co-factor Vitamin B6 metabolism 2.09 0.01195 Ascorbate metabolism, Glutathione Dehydroascorbic acid C05422 Vitamins 5.64 0.00020 metabolism

100

CHAPTER 3

Coordinated response of cyclic electron flow, photorespiration and transient starch synthesis in a high salt-evolved strain of Chlamydomonas reinhardtii

Isha Kalra, Isaiah Jacques, Xin Wang, Rachael Morgan-Kiss

101

Chapter III. Coordinated response of cyclic electron flow, photorespiration and

transient starch synthesis in a high salt-evolved strain of Chlamydomonas

reinhardtii

3.1 INTRODUCTION

In photosynthesis, linear electron flow (LEF) represents the major mechanism for transduction of light into stored energy products (ATP, NADPH), which are consumed during carbon fixation. Carbon fixation requires 3 molecules of ATP and 2 molecules NADPH; however, LEF produces an ATP:NADPH ratio of only 2.57:2, thereby leading to an overall shortage of ATP (Kramer & Evans, 2011). Plants and algae utilize several alternative pathways to compensate for this imbalance, with cyclic electron flow (CEF) around Photosystem I (PSI) being one of the major alternative pathways (Alric, 2010; Yamori & Shikanai, 2016). Although CEF accounts for less than 14% of linear electron transport during steady state conditions (Kramer & Evans, 2011), any fluctuations in the environment can trigger an increase in CEF to balance the ATP:NADPH ratio or provide photoprotection (H. Chen et al., 2015; Lucker & Kramer, 2013; Strand et al., 2015; Yamori & Shikanai, 2016). During stressed conditions, the photosynthetic electron transport chain (PETC) can become over-reduced because of bottlenecks in the downstream metabolism (Ensminger, Busch, & Huner, 2006). In such conditions, CEF can help alleviate increased excitation pressure by either increasing ATP production or by dissipating excess energy in the form of non-photochemical quenching (Munekage et al., 2002; Yamori & Shikanai, 2016). In the last decade, CEF around PSI has been shown to be important under many stress conditions when there is increased excitation pressure on the photosynthetic machinery or a need for excess ATP production (H. Chen et al., 2015; Godaux, Bailleul, Berne, & Cardol, 2015; W. Huang et al., 2010; W. Huang, Zhang, Xu, & Liu, 2017; Strand et al., 2015). The green alga Chlamydomonas reinhardtii is a great model system to study regulation of photosynthetic electron transport as a consequence of environmental changes (Hema, Senthil- Kumar, Shivakumar, Chandrasekhara Reddy, & Udayakumar, 2007). In C. reinhardtii, CEF studies have mainly focused on its role in state transition response, and short-term anoxic or extreme nitrogen stress (Iwai et al., 2010; Steinbeck et al., 2018; Takahashi et al., 2013; Terashima et al., 2012). However, only a few studies have been conducted to elucidate whether

102 there is a broader role for CEF in Chlamydomonas spp. during long-term stress acclimation (Bonente, Pippa, Castellano, Bassi, & Ballottari, 2012). To this end, a recent study by our group revealed that the high salt tolerant Antarctic Chlamydomonas sp. UWO241 displays constitutive upregulation of CEF accompanied by restructured carbon metabolism during long-term salinity acclimation (Kalra et al., 2020). Salinity stress is one of the most ubiquitous stresses imposed on photosynthetic organisms, whether they inhabit soil or aquatic environments. Soil salinity is a global issue, impacting up to 20% of soils, and has major implications for losses in crop yields (an estimated loss of 23.7 billion annually) in saline soils or under irrigation with brackish or salty water (Negrão, Schmöckel, & Tester, 2017). Improvements in crop salt tolerance could potentially mitigate salt-associated losses for future increases in crop production, expand existing cultivatable areas, and provide new opportunities for non-freshwater irrigation sources. However, plant response to salt is highly complex and improvements in crop yield traits have been generally unsuccessful (Morton et al., 2019). Genetically tractable algae, in particular green algae, have been used extensively as single-celled plant models for stress tolerance; however, at best C. reinhardtii has only a modest tolerance to salinity stress. Salinization causes osmotic stress, one of the most severe abiotic stresses for photosynthetic organisms. The individual ions are also the source of toxicity-related effects as well as nutritional imbalances. One of the main causes for losses in plant yields is downregulation of photosynthetic processes (Kumar, Wani, Penna, & Tran, 2018). Under high salinity, photosynthetic pigments, photosynthetic efficiency, and carbon fixation rates are all reduced. Salinity stress can damage photosynthetic machinery through increased oxidative stress in the form of production of reactive oxygen species (Shetty et al., 2019). In addition, green algae must continuously work to balance the osmotic pressure by active export of sodium ions or synthesis of osmoprotectant. Both these processes require excess ATP production in the cell (He et al., 2015; Kalra et al., 2020). PSI-driven CEF could contribute significantly to salinity tolerance by alleviating both oxidative stress and provide additional ATP. Currently, the role of CEF in salt tolerance has not been thoroughly reported. Salinity stress has been shown to alter downstream metabolism in many photosynthetic organisms, including enhancement of photorespiratory mechanisms (Voss, Sunil, Scheibe, & Raghavendra, 2013; Ziotti, Silva, Sershen, & Lima Neto, 2019). In plants excess salinity leads to stomatal closing and reduced CO2 uptake, promoting higher activity of the oxygenase reaction of

103

RuBisCO (Voss et al., 2013). Photorespiration metabolizes the toxic product of RuBisCO

oxygenation reaction, 2-phosphoglycolate. This mechanism leads to loss of one CO2 and one ammonium molecule. In addition, photorespiration can also use almost 50% of total ATP produced in the cell (Wingler, Lea, Quick, & Leegood, 2000). As a consequence, although photorespiration helps detox 2-phosphoglycolate, it is largely a wasteful process, depleting ATP and losing fixed carbon and nitrogen molecules. If an organism needs to evolve to be salinity resistant, constant photorespiration will exhaust ATP production and lead to losses in production. A recent study comparing drought sensitive and drought resistant plants revealed that CEF is upregulated in the drought resistant strain, however the sensitive strain upregulated photorespiration and not CEF (Lima Neto, Cerqueira, da Cunha, Ribeiro, & Silveira, 2017). More recently, Li et al. (2019) showed that photorespiratory mutants of Arabidopsis thaliana display increased rates of CEF (J. Li et al., 2019). Salinity stress imposes osmotic stress in a photosynthetic organism. To counter increased osmotic pressure, photosynthetic organisms produce osmoregulants such as glycerol and proline (Goyal, 2007a; León & Galván, 1994; Reynoso & de Gamboa, 1982). Accumulation of starch and glycerol is a common feature of salinity stressed photosynthetic organisms (Perrineau et al., 2014). Glycerol is the precursor to triacyl glycerol (TAG) biosynthesis, which are lipid droplets that are synthesized in abundance in algal cells and can serve directly as biofuel. Many algae are studied for their increased TAG production under stress conditions, especially salinity and nitrogen stress. In fact, starch to lipid biosynthesis has been shown to be a predominant mechanism for lipid accumulation in salt stressed cells of oleaginous Chlamydomonas sp. JSC4 (Ho et al., 2017). Moreover, it was shown that salinity stress is a regulator of carbon partitioning between starch and lipid, and that this mechanism is downregulated in salinity evolved strains of C. sp. JSC4 (Kato et al., 2017). Studies have also shown that salt tolerant Dunelliela salina exhibits increased carbon fixation rates and reallocation of stored starch under high salt conditions (Goyal, 2007a). However, it is still unclear whether the salt-sensitive strain C. reinhardtii can reallocate storage carbon to confer resistance to salinity during long-term acclimation. Short-term stress responses are widely studied in photosynthetic organisms, where the duration of stress lasts from minutes to hours. On the other hand, acclimation refers to exposure to stress for days to weeks and such studies have been limited. Furthermore, extensive

104 acclimation lasting from months to years leads to adaptative changes in the photosynthetic organisms that confer resistance to the stress. Although short-term salinity stress response has been widely studied and reported (Khona et al., 2016; Neelam & Subramanyam, 2013; Shetty et al., 2019; Sivakumar, Sharmila, & Pardha Saradhi, 2000; Subramanyam et al., 2010; N. Wang et al., 2018), few studies have looked at long-term salinity acclimation in the model alga, C. reinhardtii. Some of the most common changes associated with salt stress were overall reduced photosynthetic capacity, loss of light harvesting complex proteins of both PSI and PSII, loss of core PSII proteins such as CP43 and CP47 and reduced carbon fixation capacity (Neelam & Subramanyam, 2013; Subramanyam et al., 2010; N. Wang et al., 2018). Other metabolic changes, such as increased levels of oxidative stress genes, palmelloid formation and glycerol/proline synthesis were also associated with salinity stress in C. reinhardtii (Khona et al., 2016; León & Galván, 1994; Reynoso & de Gamboa, 1982; N. Wang et al., 2018). More recently, a few studies have explored the effect of long-term salinity acclimation on C. reinhardtii using transcriptome and proteome analyses, however changes associated with photosynthetic electron transport were not studied. A salinity-evolved strain of C. reinhardtii had comparable growth rates to the parent strain and had downregulated stress response genes such as oxidative stress genes (Perrineau et al., 2014). Genes involved in photosynthesis were also downregulated, however photosynthetic efficiency was not measured. Starch and glycerol biosynthetic genes were also downregulated in the evolved strain; however, they were upregulated in the parent strain during short-term salinity acclimation. Overall, it was shown that salinity evolved strain of C. reinhardtii bypassed photosynthesis and instead relied on acetate metabolism for growth and salinity resistance (Perrineau et al., 2014). A major caveat to the studies in the salt-evolved strain of C. reinhardtii is that it was adapted in mixotrophic growth media containing acetate. This is problematic because photosynthesis, which is one of the major targets for lost growth and yields, is bypassed under this scenario. The condition is also not particularly relevant for natural conditions as many unicellular green algae that belong to the genus Chlamydomonas inhabit freshwater or marine environments where acetate is not available (Shetty et al., 2019). Not surprisingly, the authors did not observe major adjustments in photosynthesis in the C. reinhardtii evolved strain. In contrast, other salt tolerant algal species exhibit significant changes in photosynthetic processes. As mentioned above, the salt tolerant Antarctic C. sp. UWO241 exhibited a restructured PSI

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complex to support constitutive CEF, (Kalra et al., 2020). To understand the full capability of C. reinhardtii to evolve better tolerance to salinity, evolution studies should be conducted in autotrophic growth media, where the organisms must rely solely on photosynthesis for growth and developing resistance to salinity. In this study, photoautotrophically grown cultures of C. reinhardtii were evolved under constant salinity stress of 100 mM NaCl for 24 months, representing a total of 125 generations. After the evolution of a salt-resistant robust growing strain of C. reinhardtii, experiments were conducted to measure growth, as well as photosynthetic and photochemical activity. Furthermore, to understand the metabolic changes associated with salinity evolution, whole cell proteome of the high salt grown evolved strain and the parent strain were analyzed and compared to the proteome of the parent strain grown in low salinity. We hypothesized that higher rates of CEF plays a role long-term salinity tolerance in C. reinhardtii. To the best of our knowledge, this is the first study attempting to elucidate the role of CEF in long-term salinity acclimation and the associated metabolic changes in a photoautotrophic salt resistant C. reinhardtii strain.

3.2 METHODS

3.2.1 Growth conditions and evolution of high salt evolved strain of C. reinhardtii

Chlamydomonas reinhardtii UTEX 89 (Wildtype, abbreviated as Wt henceforth) was grown in Bold’s basal media (BBM) (Nicholas and Bold, 1965) and maintained in a photo-incubator at

23°C in 250 ml flasks. The cultures were grown under ambient CO2 conditions and constant light regime (100 μmol photons m-2s-1.) on a rotatory shaker. For the evolution of a high salinity evolved strain (Evolved strain, Ev henceforth) of C. reinhardtii, a single colony from the parent C. reinhardtii (Wt) strain was inoculated into BBM (low salt, LS) and then transferred to BBM supplemented with 100 mM NaCl (High salt, HS). Serial transfers were made in HS media every two weeks with 10% of inoculum for 24 months and cultures were maintained as described above.

3.2.2 Experimental set-up

For the salinity acclimation vs adaptation experiment, cultures were grown in temperature regulated aquaria in 250 ml pyrex tubes with temperature/light regime of 23°C /100 μmol photons m-2s-1. Cultures were continuously aerated using sterile ambient air from air pumps

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(Morgan et al., 1998). The stock cultures of parent and evolved strain were grown in LS and HS media in the aquaria to acclimate in the new environment. Exponentially grown cells of the Wt and Ev were inoculated into both LS and HS conditions giving rise to four different growth conditions: Wt-LS, Wt-HS, Ev-LS, Ev-HS. Briefly, the stock cultures of Wt and Ev strain were centrifuged at 1000xg for 5 min and the medium was removed. The cells were then washed in the respective new media (LS or HS) 3 times to get rid of any previous media residues. Three biological replicates for each condition were set up in pyrex tubes in the aquarium as described above. Optical density (750 nm) was measured along the growth curve using a spectrophotometer and cells/ml were calculated using Countess cell counter (Thermo Fisher, Cat no. A27977) to assess growth rate. Chlorophyll was extracted from whole cells using 90% acetone and bead-beating to break the cells. Absorption was measured at 647 and 664 nm, and chlorophyll a and b were calculated according to Jeffry and Humphrey (Jeffrey & Humphrey, 1975)

3.2.3 PSII measurements and oxygen evolution rates

Photosynthetic measurements for PSII were done using Dual-PAM 100 (Walz, Germany). Briefly, 2 ml of culture suspension were far red adapted for 2 min in presence of 20 μl of 0.5 M

NaHCO3 prior to the measurement. Minimal fluorescence (FO) was measured for dark-adapted cells, while maximal fluorescence FM was measured in the dark-adapted state after a saturation pulse of 10,000 μmol photons m-2s-1. Cultures were measured for maximum photosynthetic efficiency (FV/FM), PSII yield (YPSII), non-photochemical quenching (NPQ), non-regulated non-photochemical quenching (NO) and fraction of open PSII centers (qL) throughout the growth curve using the induction curve with actinic light intensity of 93 μmol photons m-2s- 1. Light curves were also measured on mid-log phase cultures to identify the potential for NPQ and establish saturating light intensity. Oxygen evolution rates were measured on mid-log phase cultures using Clark-type oxygen electrode (Hansatech Instruments) after brief incubation in the

dark. Whole-cell oxygen evolution was measured for 2 min in the presence of 20 mM NaHCO3 and final rates were normalized to cell density.

3.2.4 P700 photo-oxidation and cyclic electron flow

P700 oxidation-reduction activity was measured as absorbance changes at 820 nm using a Dual PAM-100 instrument (Walz, Germany). Exponential phase cultures equivalent to 25 ug

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chlorophyll were dark adapted for 10 min and filtered on GF/C filters (Whatman). Using the leaf attachment on the Dual-PAM, P700 photo-oxidation were measured using far red preferential photooxidation of P700 as described in Morgan et al, 2002 (R. M. Morgan-Kiss et al., 2002). Prior to the measurement the signal was balanced and measuring light was switched on. Far-red light (λmax=715 nm, 10 Wm−2, Scott filter RG 715) was then switched on to oxidize P700 and the proportion of photooxidizable P700 was calculated as ∆A820/A820. After achieving steady state oxidation level, the FR light was switched off to re-reduce P700. The half-time of re-

+ red reduction of P700 to P700 ( t½ ) was used as a relative measure of PSI driven CEF and calculated using first-order exponential-decay kinetics as described before (Ivanov, Morgan, Gray, Velitchkova, & Huner, 1998). Calculations and statistical analysis were done using Prism software (n=9).

3.2.5 Protein extraction and sample preparation for proteomics

Mid log phase cultures (25 ml) were collected for each strain and condition by centrifugation at

2500g for 5 min. Cell pellets were flash frozen in liq N2 and stored at -80°C until use. Whole cell proteins were extracted as described before with some modifications (Valledor, Luis; Weckwerth, 2014). Briefly, cells pellets were resuspended using extraction buffer (100 mM Tris (pH 8.5), 10% glycerol (v/v), 10 mM Dithiothretol (DTT), 1.2% (v/v) plant protease inhibitor cocktail (Sigma, Cat P9599), 2 mM Pefabloc and 20 mM sodium fluoride (NaF). The resuspension was transferred to a screw cap tube containing 50 mg silica beads (Cat. No. A6758, Biorad) and homogenized using a bead beater (BioSpec) for 1.5 min (10 s ON, 2 min OFF, 9x). SDS solution (20% w/v) was added to the homogenized cell suspension and mixed gently via inversion, followed by incubation at 95°C for 5 min. The cell suspension was then centrifuged at 12,000g for 30 min to pellet any insoluble material. Protein concentration was calculated using Bradford method and 100 ug of protein was aliquoted for further processing for proteomics. The aliquoted protein was precipitated using acetone/βmercaptoethanol (0.5%) solution and incubated at -20°C overnight. The precipitated protein sample was centrifuged, and the pellet was washed in the acetone/βmercaptoethanol (0.5%) solution. This was followed by another wash with 100% acetone followed by 90% acetone to remove any impurities. Finally, the protein pellet was air dried and was processed for proteomics as described in (X. Wang et al., 2016). Briefly, the protein pellet was resuspended in denaturing buffer containing Tris-HCl (50 mM, pH

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8.0), 8 M Urea, 5 mM DTT and incubated at 37°C for 1 hour. This was followed by alkylation with 15 mM iodoacetamide (IAA) for 30 min in dark at room temperature. The protein samples were diluted 4 fold and digested with Mass-spectrometry Grade Trypsin Gold (Promega, Madison, WI) at 1:100 w/w concentration for 18 hours at 37°C with gentle rotation. Sep-Pak C18 plus desalting columns (Waters Corporation, Milford, MA) were used to clean up the digested samples.

3.2.6 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS)

Proteomics analyses was done as previously described (Shinde et al., 2020) using Thermo LTQ Orbitrap XL mass spectrometer. Briefly, the full mass spectra were recorded at 350-1800 m/z at a resolution of 30,000, and the 12 most abundant peaks of each scan were selected for MS/MS analysis. Thermo Scientific MSFilereader was used to directly work with the.RAW data files. Proteomic analysis was done using Patternlab for Proteomics 4.0 software following published guidelines (Carvalho et al., 2016). Briefly, C. reinhardtii protein database was downloaded from NCBI containing both Swiss-Prot and TrEMBL entries. Subsequently a target-decoy database was generated using reverse sequences and including 127 common proteomic contaminants. A Comet PSM search was initiated using the above database and the sample sequences. The results were filtered using SEPro and differential protein expression was identified using T-Fold test with false discovery rate set to 0.05.

3.3 RESULTS

3.3.1 The high salinity evolved strain has faster growth rate

Following 104 weeks of serial transfer in the high salt media (HS, BBM + 100 mM NaCl) to create the evolved strain, comparison of growth rates between Wt and Ev strains was performed. The C. reinhardtii wild type strain under low salinity (Wt-LS) displayed a typical growth curve (Fig. 3.1, circles) under photoautotrophic conditions, with a growth rate of 0.0088±0.008 hr-1 (Table 3.1). However, under high salinity (100 mM NaCl), growth of the wild-type strain (Wt- HS) was impaired (Fig. 3.1), reaching the stationary phase faster and had a significantly lower (p=0.029, t-test) growth rate of 0.0057±0.0010 hr-1, relative to Wt-LS cultures (Fig. 3.1, squares, Table 3.1). On the other hand, under long-term salinity stress the evolved strain (Ev-HS) showed robust growth and had a similar growth (ns, t-test) rate as the wildtype strain under low salinity

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(0.0083±0.004 hr-1, Fig. 3.1, triangles, Table 3.1) and 35% (p=0.027) higher growth rate than Wt-HS. We also tested the growth rate of the evolved strain under low salinity (Ev-LS) condition to see if there are any changes associated with growth of a high salinity evolved strain when the stress is eliminated. Remarkedly, the evolved strain under low salinity displayed a 22.7% higher growth rate (p=0.016) compared with Wt grown under comparable conditions (0.0104±0.004 hr- 1, Fig. 3.1, Table 3.1).

3.3.2 The evolved strain maintains high photosynthetic capacity and constitutive upregulated NPQ under low and high salinity

PSII light curves were conducted for wild-type and evolved strains using dual PAM and actinic light intensities of the range 20-800 μmol photons m-2s-1 (Fig. 3.2). Under both low and high salinity, the evolved strain maintained higher YPSII than either Wt-LS or -HS (Fig. 3.2 A). The proportion of open PSII centers (qL) was lowest for Wt-HS under all light intensities, whereas Wt-LS had comparable qL values to both the Ev-LS and Ev-HS (Fig. 3.2 B). The proportional non-photochemical quenching (YNPQ) was markedly higher for Wt-LS and both Ev-LS and -HS as compared to Wt-HS (Fig. 3.2 C). Interestingly, the absolute yield of non-photochemical quenching (NPQ) was significantly higher for both Ev-LS and Ev-HS with increasing light intensities than both Wt-LS and Wt-HS (Fig. 3.2 D). Wt-HS strain had the least capacity for NPQ induction. On the other hand, Wt-HS displayed the highest capacity for nonregulated energy dissipation (YNO) under increasing light regime (Fig. 3.2 E). Both the evolved strain conditions had a decrease in YNO capacity as the light intensity increased. Thus, the evolved strain maintains high capacity for NPQ under both low and high salinity, whereas the wildtype switches from high NPQ capacity under low salinity to high NO capacity under high salinity. To further investigate the relationship between strain-specific differences in induction of YNPQ and YNO under increasing light intensity, as both are energy dissipative processes albeit with different pathways, we compared trends of both processes between Wt-HS and Ev-HS (Fig. 3.2 F). With the increasing light intensity Ev-HS displayed no change in NO capacity but induced significantly higher NPQ response, which was 1.4 folds higher than Wt-HS at the maximum light intensity. On the other hand, Wt-HS increased both NPQ and NO response to dissipate excess energy due to increased excitation pressure at higher light intensities.

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3.3.3 The evolved strain has higher oxygen evolution and respiration rates compared to the parent strain

We compared both oxygen evolution and dark respiration rates for Wt and Ev strains grown under low and high salinity respectively (Fig. 3.3). The evolved strain showed a significantly higher (1.5-fold, p<0.01) oxygen evolution rates under high salinity as compared to wildtype strain under low salinity (Fig. 3.3 A). The evolved strain also showed higher dark respiration rate as compared to wild type under low salinity (Fig. 3.3 B).

3.3.4 The evolved strain displays constitutive high rates of PSI-CEF

P700 photo-oxidation/reduction kinetics were measured to understand the effect of long-term

+ adaptation vs acclimation to salinity. The change in absorbance of P700 radical after far-red preferential PSI-excitation was measured as ∆A820 /A820 (Fig. 3.4 A). The Wt-LS strain had the largest relative level of FR-oxidized P700 (mean ∆A820 /A820 = 0.31). Wt-HS (mean ∆A820 /A820 = 0.17) displayed significantly lower (~1.8 folds) ∆A820 /A820 compared to Wt- LS. Both Wt-HS and Ev-HS (mean ∆A820 /A820 = 0.17 and 0.093 respectively), had significantly lower values compared to Wt-LS, indicating that PSI reaction center was impacted by salinity, regardless of the acclimation/adaptation time; however, Wt-HS exhibited 1.8 folds higher ∆A820 /A820 relative to Ev-HS. Last, Ev-LS displayed 1.5 folds higher ∆A820 /A820 than Ev-HS, indicating that removing the stress restores some P700 activity in the evolved strain.

red The dark re-reduction rate (t½ ) of P700 was measured post FR illumination (Fig. 3.4 B). Under FR illumination when only PSI received excitation energy, alternative electron transport pathways, mainly PSI-CEF, are responsible for re-reduction of PSI (Ravanel et al, 1994, Ivanov red red et al, 2012). Ev-HS had significantly faster t½ (mean t½ = 335.4 ms) compared to other three conditions, signifying that the Ev strain grown in high salt possessed the highest rates of CEF. red Interestingly, Ev-LS (mean t½ = 579.6 ms) had 1.3 folds faster CEF compared to Wt-LS (mean red t½ = 745.4 ms). Thus, the evolved strain shows constitutive upregulation of CEF rates. Wt-HS (mean t½red = 473.7 ms) also maintained 1.6 folds faster t½red as compared to the parent wildtype strain (Wt-LS). Thus, high salinity induced faster CEF in both the Wt and Ev strains; however, CEF was further enhanced in the Ev strain.

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3.3.5 Proteomic comparison of the evolved and the wild type strains under high salinity

We hypothesized that alterations in the PETC in response to high salinity might be reflected in differential expression of proteins in the Wt vs. Ev strains. To understand the changes in the protein profile of C. reinhardtii associated with short-term (6 days) and long-term (24 months) salinity acclimation response, whole cell proteomes of Wt-LS, Wt-HS and Ev-HS were assessed. Cellular proteins from three biological replicates of each strain were extracted and the peptide sequences were matched with C. reinhardtii proteome database (Figs 3.5 & 3.6). The proteomes of Wt-HS and Ev-HS were individually compared with Wt-LS and a list of significantly differentially regulated proteins was generated using TFold analysis (Pattern Lab 4.0) with the filtering parameters q-value 0.05, F-stringency 0.04 and L-stringency 0.60.

3.3.5.1 Sub-cellular localization

The significantly differentially regulated proteins were analyzed and categorized into their predicted sub-cellular localization (Fig 3.6). Under high salinity conditions, the wild type and evolved strain had 100 and 124 significantly differentially regulated proteins, respectively. (Fig 3.5), out of which they share 64 common proteins (Fig 3.6 A). For each sub-cellular categorization, the number of proteins were differentiated into up- and down-regulated (Fig. 3.6 B). In Wt-HS cultures, the maximum number of proteins were localized in the chloroplast (21), with ~60% upregulated and 40% downregulated. On the other hand, although the evolved strain also has the highest number of proteins localized to the chloroplast (22), most of the proteins are downregulated (~ 86 %) in response to high salinity (Fig 3.6). The cytoplasm was the site with the second-highest number of proteins localized for both Wt-HS (16) and Ev-HS (18); however, Wt-HS has ~ 60% proteins upregulated, while Ev-HS had almost 90% proteins upregulated. Twelve mitochondrial proteins were differentially expressed in Wt-HS while fourteen proteins were differentially regulated in Ev-HS. Furthermore, 8 proteins in Wt-HS and 10 proteins in Ev- HS were localized in the nucleus. However, most nuclear proteins (90%) were downregulated in Wt-HS as compared to only 20% in Ev-HS. Last, only a few proteins (3 in Wt-HS and 2 in Ev- HS) are located in the ER and all were upregulated.

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3.3.5.2 Gene ontology

Next, Gene ontology molecular functions were assigned to each significantly differentiated protein of Wt-HS and Ev-HS (Fig. 3.7). Most of the proteins in Wt-HS and Ev-HS belong to the class of oxidoreductases. The second most common class of proteins in Wt-HS are transferases (17 proteins), out of which 7 are upregulated and 10 are downregulated. On the other hand, RNA binding proteins are the second most common functional category in Ev-HS (total 13 proteins), followed by transferases (total 8 proteins). Interestingly, DNA binding proteins are mostly upregulated (~80%) in Ev-HS as compared to Wt-HS, where all are downregulated.

3.3.5.2 Kegg Orthology Biological Pathway

To understand the changes in the metabolic pathways under short-term and long-term salinity acclimation, significantly regulated proteins of Wt-HS and Ev-HS were categorized into Kegg Orthology (KO) biological pathway (Fig. 3.8). The significantly regulated proteome of Wt-HS and Ev-HS compared to Wt-LS, contained many identical proteins that the two strains shared under the high salinity conditions, informing us of a set of core proteins that are important in both short-term and long-term salinity acclimation. There were 64 common proteins that the two strains shared out of which 27 are upregulated and 37 are downregulated (Fig. 3.9 A and B. respectively). On the other hand, there were also a total of 60 uniquely differentially regulated proteins of Ev-HS (Fig. 3.10 A) and 40 unique proteins in Wt-HS (Fig. 3.10 B).

Cytochrome b6f, CBB enzymes, carbon-dioxide uptake and photorespiration are downregulated in the evolved strain after long-term salinity acclimation

As the Chlamydomonas strains were grown in strict photoautotrophic conditions, photosynthesis provided the only fixed carbon source for the cells. Both photosynthetic electron transport (PET) and carbon fixation (CBB cycle) were major sites for changes to the proteome under both short- term and long-term salinity acclimation. During short-term acclimation (ie. Wt-HS samples), 10% of the significantly regulated proteins belonged to PET (7 proteins) and CBB (3 proteins) together. The light harvesting complex proteins of PSII (lhcb3 and 5) and PSI (lhca3) were both upregulated in Wt-HS (Table 3.2). The PSI core protein (PsaD) and proteins of oxygen evolving complex of PSII (PsbO and Q) were also upregulated. On the other hand, three subunits (PETA, PETC and PETO) of cytochrome b6f were downregulated. In the CBB cycle, fructose-1,6-

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bisphosphatase and fructose-1,6-bisphosphate aldolase (FBA) were both upregulated; however, rubisco activase was downregulated. FBA have been shown to be important in abiotic stress response, including salinity (Lu et al., 2012). Interestingly, the oxidative pentose phosphate pathway (OPPP) protein 6-phosphoglucanate dehydrogenase (PGDH) was upregulated by 3 folds. PGDH is responsible for decarboxylation of 6PG to generate Ru5P. Under long-term salinity stress (ie. Ev-HS conditions), 10% of the total proteins significantly regulated belonged to PET (6 proteins) and CBB cycle (7 proteins). The light harvesting complex (LHC) proteins of PSII (lhcb3 and lhcb5) were upregulated but core protein

(PsaC) and LHC protein of PSI were downregulated. Similar to Wt-HS, two subunits of cyt b6f (PETC and PETO) were downregulated in Ev-HS (Table 3.3). Carbon fixation proteins were predominantly downregulated, with the exception of FBPase which was upregulated under high salinity. Several proteins involved in regeneration of ribulose-5-phosphate (Ru5P): phosphoglycerate kinase (PGK), sedoheptulose-bis-phosphatase (SBPase) and Ribulose 5- phosphate isomerase (RPI) were downregulated in Ev-HS. Interestingly, similar Wt-HS, the OPPP protein 6-phosphogluconate dehydrogenase (PGDH), responsible for regenerating Ru5P from 6-phosphogluconate was upregulated by 2.5-fold. Furthermore, both RubisCO small subunits (RbcS1 and 2) were downregulated. RbcS have been shown to degrade under oxidative stress conditions (Knopf and Shapira, 2005). Finally, rubisco activase was also downregulated, similar to Wt-HS. Carbon concentrating mechanisms (CCM) operate in C. reinhardtii under conditions of

low CO2, increasing the CO2:O2 ratio around RuBisCO to increase carbon fixation capacity. This - can occur in several ways: active transport of HCO3 across membrane, inorganic carbon

conversion via carbonic anhydrases (CA) and active CO2 uptake. There are several proteins that

have been shown to be important in active uptake of CO2 (Low CO2 inducible (LCI) proteins: LCIB, LCIC, LCID and LCIE) (Wang et al, 2015). In Wt-HS, two LCI proteins, LCIC and LCIE were significantly downregulated. Another protein, peptidylprolyl isomerase (FKB19) was also downregulated. Incidentally, cyclophilin proteins that contain peptidylprolyl isomerase domain have been shown to be involved in low CO2 acclimation response in C. reinhardtii (Somanchi

and Moroney, 1999). Several LCI proteins involved in CO2 uptake were significantly downregulated (LCIC, LCID and LCIE) in Ev-HS. Interestingly, two cyclophilin type PPI were also downregulated under long-term salinity. In algal species, glycolate dehydrogenase (GYD) is

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an important photorespiration protein, responsible for oxygenation of glycolate to produce

glyoxylate and hydrogen peroxide (H2O2). We found that both under short-term and long-term salinity acclimation conditions, GYD was significantly downregulated. However, the fold change under long-term acclimation was higher (3.5 folds) as compared to short-term acclimation (- 2.66 fold). We also found that catalase (CAT) was also downregulated in Ev-HS.

CAT is responsible for scavenging H2O2 generated by the above reaction. Together, downregulation of GYD and CAT in the evolved strain demonstrates a downregulation of photorespiratory response. The combined downregulation of CCM proteins involved in carbon

dioxide uptake and photorespiration suggest a possible high CO2/low O2 environment in the evolved strain.

Mitochondrial TCA cycle and oxidative phosphorylation are upregulated in the evolved strain after long-term salinity acclimation Pyruvate metabolism was downregulated under short-term salinity acclimation: pyruvate carboxylase (PYC) and pyruvate kinase (PK) were both downregulated. PYC catalyzes the irreversible reaction to form 4 carbon oxaloacetate from 3 carbon pyruvate, utilizing 1 molecule

of CO2 and ATP. On the other hand, PYK catalyzes the reversible reaction between oxaloacetate and phosphoenol-pyruvate (PEP). The combined downregulation of PYC and PK suggest accumulation of pyruvate in the cell, which may be used for other downstream reactions. On the other hand, the mitochondrial TCA cycle was upregulated under short-term salinity acclimation, with isocitrate dehydrogenase (IDH) and malate dehydrogenase (MDH) showing significant upregulation. After long-term salinity acclimation, pyruvate metabolism was downregulated. Phosphoglycerate kinase (PGK) and phosphopyruvate hydratase (PH) that catalyze the reversible reaction of glyceraldehyde-3-phosphate (G3P) to PEP were downregulated in Ev-HS. Additionally, pyruvate dehydrogenase (PDH) E2 enzyme, which catalyzes the irreversible conversion of pyruvate to acetyl-CoA was also significantly downregulated. On the other hand, TCA cycle was upregulated after long-term salinity acclimation. IDH, MDH and aconitate hydratase (ACH) enzymes of the mitochondrial TCA cycle were significantly upregulated in Ev- HS. Oxidative phosphorylation was also upregulated in evolved strain but not in the wildtype strain during salinity acclimation. Three proteins of mitochondrial respiration were upregulated

115 in the Ev-HS: cytochrome oxidase subunit 2 (cox2b), ATP synthase associated proteins 1 and 3 (ASA1 and ASA3). Interestingly, it has been previously reported that salinity tolerant plants display altered mitochondrial proteins with increase in MDH, ADH and alternate oxidase (Jacoby et al., 2013).

Transient starch synthesis and breakdown, as well as ethanol and glycerol biosynthesis are all upregulated in the evolved strain Proteins involved in gluconeogenesis and glycolysis were significantly upregulated under short- term and long-term salinity acclimation. FBPase, FBA and phosphoglucomutase (PGM) were upregulated in Wt-HS while FBPase and PGM were upregulated in Ev-HS. Starch metabolism was upregulated under both short-term and long-term salinity acclimation (Fig. 3.8, Tables 3.2 and 3.3). Chloroplastic starch synthase (STA2) was upregulated for both strains but all other proteins are involved in starch breakdown (Alpha-amylase, Alpha-1,4-glucan phosphorylase) and display higher fold change than STA2. Glycerol-3-Phosphate dehydrogenase (GPD) is an important protein in glycerol and lipid biosynthetic pathways. Three different isoforms of GPD were upregulated in both Wt-HS and Ev-HS, but the fold change of the isoforms in Wt-HS was 1.5 times higher than Ev-HS. Recently it was shown that an oleaginous alga Chlamydomonas sp. JSC4 switches from starch-to-lipid biosynthesis under salinity conditions (Shih-Hsin Ho et al., 2017). On the other hand, fatty acid biosynthesis was downregulated in Wt-HS. Three proteins belonging to FA biosynthesis: biotin carboxylase, CoA carboxyltransferase N-terminal domain-containing protein and 3-oxoacyl-[acyl-carrier-protein] synthase were downregulated in Wt-HS. Biotin carboxylase and CoA carboxyltransferase N-terminal domain-containing protein are part of the Acetyl-coA carboxylase (ACC) that converts acetyl-CoA to malonyl-CoA, while 3-oxoacyl-[acyl-carrier- protein] synthase is involved in the condensation of fatty acid biosynthesis. On the other hand, polyunsaturated fatty acid biosynthesis was upregulated in the evolved strain after long-term salinity acclimation, via upregulation of cytochrome b5 reductase protein (Wayne et al., 2013). However, four other proteins involved in fatty acid biosynthesis are down-regulated including biotin carboxylase, 3-oxoacyl-reductase and CoA carboxyltransferase of the ACC complex as well as PKS_AT domain-containing protein. Alcohol/aldehyde dehydrogenase (ADHE) is a bifunctional enzyme that is involved in fermentation and converts acetaldehyde to ethanol. In the

116 chloroplast, pyruvate can either be used for regeneration of oxaloacetate (OA) through pyruvate carboxylase or it can be used for fermentation. In our proteome analysis, we found that ADHE is significantly upregulated in both Wt-HS and Ev-HS with similar fold change (3.86- and 3.55- fold, respectively). Recently, it has been shown that starch accumulates concomitantly with increases in ADHE levels, perhaps because algae use internal stored starch to divert towards fermentation (van Lis et al., 2017). Increased abundance of ADHE have also been shown under anoxia, zinc deficiency and in CCM deficient strains (van Lis et al, 2017). Last, nucleotide sugar metabolism was upregulated in both the strains. Mainly three core proteins involved in conversion of glucose-1-phosphate to UDP-D-xylose and UDP-D-arabinose were upregulated (Reiter, 2008; Yin et al. 2011).

Reactive oxygen and nitric oxide scavenging proteins are differentially regulated in the evolved strain Many environmental stresses, including salinity stress, lead to redox imbalance and subsequently the formation of reactive oxygen species (ROS, oxidative stress) and reactive nitrogen species such as nitric oxide (NO) (Hancock et al, 2002). Photosynthetic organisms balance redox poise by upregulating ROS and NO scavenging pathways. In our proteome, we observed two proteins involved in nitric oxide signaling to be significantly regulated in both Wt-HS and Ev-HS. Cytochrome P450 (nitric oxide reductase, NOR) was highly upregulated while the nitrogen regulatory protein PII (GLB1) was downregulated. GLB1 transcript levels have been shown to be negatively regulated by NO (Zalutskaya et al., 2018). Proteins involved in oxidative stress pathways were also differentially regulated in both the conditions. Wt-HS and Ev-HS both upregulated mitochondrial superoxide dismutase under high salinity, however in Wt-HS the fold change was higher. Interestingly, two other antioxidative enzymes, L-ascorbate peroxidase and catalase were both downregulated only in Ev-HS.

The evolved strain differentially regulates protein folding and translation

Salinity stress changes the physiological environment of cells and can lead to changes in protein folding processes. Salinity stress has been shown to play a role in unfolded protein response (UPR) that occurs following exposure to stress, when unfolded proteins are accumulated in the ER (Liu et al, 2007). In our proteome we found several chaperone proteins that were

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differentially regulated in both Wt-HS and Ev-HS (Fig. 3.8, Tables 3.2 and 3.3). In Wt-HS, 4 proteins in this category were upregulated and 3 were downregulated. Protein disulfide isomerase (PDI2 and PDI) and calreticulin proteins were upregulated whereas chaperonin 11 and GrpE proteins were downregulated. On the other hand, in Ev-HS, a different set of chaperones were involved in the salinity response. Protein disulfide isomerase, peptidyl prolyl isomerase (FKB12), heat shock protein 90A and chaperonin 60 were upregulated while chaperonin 11, GrpE protein and a FKPB type peptidyl prolyl isomerase was downregulated. Recently it was shown that accumulation of FKB12 transcript in Arabidopsis increases resistance to multiple stresses (Alavalli et al, 2018). Also, chaperonin 60 has been shown to be crucial for Rubisco’s assembly in Arabidopsis (Aigner et al, 2017). Amino acid metabolism was differentially regulated after long-term salinity acclimation in the evolved strain. Two proteins involved in lysine biosynthesis along with SHMT protein of glycine, serine and threonine metabolism were downregulated but METM protein involved in cysteine and methionine biosynthesis was upregulated (Fig. 3.8, Tables 3.2 and 3.3). Translation machinery was also significantly regulated in Ev-HS with 7 proteins downregulated while 11 were upregulated. Majority of the upregulated translational proteins were localized to the cytoplasm while the down-regulated proteins were predominantly localized to the chloroplast and mitochondria. On the other hand, fewer proteins (total 8) belonging to the translation machinery were significantly regulated in Wt-HS, out of which 2 were upregulated and 6 were downregulated.

3.4 DISCUSSION

In this study, we investigated the impact of adaptation of the salt-sensitive alga C. reinhardtii to a higher salinity of 100 mM NaCl conditions. While previous researchers have performed similar evolution studies on C. reinhardtii, to our knowledge this study is one of the first to use autotrophic conditions. In contrast, most previous studies included an organic carbon source, acetate, in the growth medium (ie. TAP medium). After 24 months of evolution representing 125 generations, the evolved strain of C. reinhardtii grew significantly better than the parent strain under high salinity conditions and had a comparable growth physiology under high salinity as Wt grown under low salinity (Fig. 3.1, Table 3.1). Moreover, the Ev strain exhibits the ability to grow faster and achieve higher biomass

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yields compared to the Wt strain when the high salt stress was alleviated. Higher growth rates in the Ev strain were accompanied by higher rates of photosynthesis. Our results are in contrast with previous studies that reported reduced growth and photosynthesis in Chlamydomonas strains which were evolved in high salt-TAP medium (Sithtisarn et al. 2017). How does the evolved strain achieve higher growth rates? To answer this question, we probed the photochemical activity of PSI and PSII in parallel with comparative whole cell proteomics to understand changes in photosynthetic, metabolic and stress proteins associated with long-term salinity acclimation in the evolved strain. Several studies have reported that photosynthesis is significantly hampered by salinity stress. Salinity stress has been shown to decrease photosynthetic capacity (Allakhverdiev & Murata, 2008; Subramanyam et al., 2010) and shows down-regulated levels of several photosynthetic proteins including PSII and PSI core proteins, LHCI and II proteins (Neale & Melis, 1989; Shetty et al., 2019). During salinity acclimation, Wt had significantly lower photosynthetic yield (YPSII) and proportion of open PSII (qL) (Appendix Fig. 3. 1 B, C), while Ev maintained high FV/FM, YPSII and qL throughout growth (Appendix Fig. 3. 1), as well as a higher capacity to maintain higher YPSII levels under increasing actinic light (Fig. 2 A). Whole cell proteomic analyses revealed changes in major photosynthetic proteins in both Wt and Ev strains. Several LHCII subunits were upregulated under both short-term (Wt-HS) and long-term

(Ev-HS) salinity acclimation, while cyt b6f subunits were downregulated under both conditions. We also observed that CEF levels were upregulated in the Wt-HS strain, suggesting that CEF is important for short-term salinity acclimation. However, the Ev strain displayed faster rates of CEF under low salinity conditions which was further enhanced when the strain was grown in high salt (Fig. 4). This “constitutive CEF” phenotype has been previously observed in the halotolerant Antarctic alga Chlamydomonas sp. UWO241 which also exhibits higher CEF rates regardless of the salinity levels (Kalra et al., 2020). During CEF electrons are cycled around PSI, cyt b6f and plastocyanin and back to PSI. This leads to formation of proton gradient across cyt b6f, that can either lead to increased ATP production or NPQ response via acidification of the thylakoid lumen (Lucker & Kramer, 2013). Thus, the increased CEF in the Ev strain may help in increased ATP production to be used in the downstream metabolic reactions for maintaining growth and osmolarity and/or used for initiating NPQ response.

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NPQ is induced as a photoprotection mechanism when the photosynthetic machinery is exposed to high excitation pressure, for example ether high light stress or under any condition where the plastoquinone pool is over-reduced (Erickson, Wakao, & Niyogi, 2015). Under conditions of high excitation pressure, the excess electrons in the photosynthetic electron transport chain (PETC) can be diverted to oxygen molecules, generating reactive oxygen species (ROS) ((Roach & Krieger-Liszkay, 2014)). NPQ-induced downregulation of PSII represents one mechanism for reducing ROS production. In the current study, the Ev strain exhibited a higher capacity for NPQ under either low or high salinity conditions, relative to the Wt strain (Fig. 2 C, D). In addition, Ev strains also displayed higher steady-state NPQ capacity during early and late- log growth phase (Appendix Fig 3.1 D). In C. reinhardtii, LHCSR (Light Harvesting Complex Stress related protein) proteins 1 and 3 are crucial for initiating NPQ response (Peers et al., 2009); however, we did not detect differential expression of LHCSR proteins in the Ev proteomes. This was not particularly surprising, since LHCSR proteins are expressed during very short-term high light treatments (Nawrocki, Liu, & Croce, 2020), and therefore may not play a role in longer term NPQ responses such as those observed in this study. We propose that constitutive increase in CEF in the evolved strain primes the organism to increase NPQ response to dissipate excess energy under high light conditions. In contrast with the Ev strain, despite higher CEF rates in Wt-HS conditions, NPQ capacity was not affected in the Wt-HS. One possible reason for this is the downregulation of phototropin protein in the Wt strain (Fig. 3.10 B), that plays a role in NPQ response (Petroutsos et al., 2016). The question arises, what is the use of CEF upregulation in short-term salinity acclimation? One possibility is that increased CEF in the Wt is primarily used for increased ATP production which could be used for downstream metabolic reactions to help balance osmotic pressure induced by salinity. CEF is also known to balance the ATP:NADPH demand for carbon fixation (Kramer & Evans, 2011). In a recent study, Kalra et al. (2020) showed that under high salt (700 mM NaCl) growth conditions, constitutive CEF was correlated with upregulation of many key CBB cycle enzymes in the halotolerant psychrophile, Chlamydomonas sp. UWO241. Although the evolved strain also exhibited high CEF, unlike the Antarctic strain, several CBB cycle enzymes were downregulated after long-term salinity acclimation with the exception of FBPase, which was upregulated (Table 3.3). Notably, enzymes participating in Ru5P regeneration were downregulated, along with reduced content of RuBisCO small subunit and Rubisco activase.

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RuBisCOs have been shown to degraded under oxidative stress conditions (Knopf & Shapira, 2005). The downregulation of several key CBB cycle enzyme in the Ev-HS is somewhat surprising, since it exhibits the ability to maintain comparable growth as Wt-LS. As discussed below, we suggest that additional changes to downstream metabolism in the Ev strain allow it to maintain robust growth even when the Calvin cycle is downregulated.

The RuBisCO enzyme catalyzes two reactions, incorporating either CO2 or O2 as substrates, and leading to very different outcomes. While carboxylation of RuBP produces two molecules of 3-phosphoglycerate (3PGA), the oxygenation reaction of RuBisCO produces only

one molecule of PGA and one molecule of 2-phosphoglycolate (2PG). Photorespiration (PR) scavenges 2PG by oxidizing glycolate to produce glyoxylate via the enzyme glycolate dehydrogenase. Unlike plants, where photorespiration occurs in peroxisomes via glycolate oxidase, algae possess only mitochondrial glycolate dehydrogenase. The detoxification of glyoxylate occurs in a series of reactions, ultimately resulting in the loss of carbon dioxide and ammonia (Fang et al., 2012). Photorespiration is active whenever a condition leads to higher RuBisCO oxygenase activity, and in plants plays an important role in abiotic stress response. Photorespiration is also a sink for ATP, utilizing up to 50% of ATP produced in the cell (Wingler et al., 2000), and is thought to dissipate excess energy and reducing equivalents under a variety of stresses (Voss et al., 2013). In our Ev-HS proteome, several proteins of the mitochondrial photorespiratory machinery were downregulated after long-term stress acclimation (GDH, HPR1, SHMT and GSCP) (Table 3.3). All of these proteins have been shown to be significantly upregulated under conditions of low CO2, a major trigger of PR activation (Fang et. al., 2012). In addition, several proteins of the CCM (Y. Wang, Stessman, & Spalding, 2015) participating in

CO2 uptake (LCIC, LCID and LCIE) were also downregulated, suggesting a possible high

CO2:low O2 environment in the chloroplast. Although glyoxylate dehydrogenase is widely implicated in conversion of glycolate to glyoxylate (Aboelmy & Peterhansel, 2014) some studies have shown it to play a minor role in mitochondrial photorespiration algae such as Dunaliella or C.reinhardtii (Goyal, 2007a; Goyal & Tolbert, 1996). A light activated glycolate to glyoxylate conversion occurs in the chloroplast of algae, utilizing CEF. The continuous cycling of glycolate to glyoxylate would lead to increased ATP production but no NADPH. Additional ATP was shown to be important for bicarbonate transport into the chloroplast to increase CO2 levels (Goyal & Tolbert, 1990). As the evolved strain maintains high rates of CEF in combination with

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lowered levels of PR and LCI proteins, we suggest that the evolved strain sustains high growth yield through downregulation of wasteful mitochondrial photorespiration via increased CEF- mediated glycolate oxidation in the thylakoid membrane. We also hypothesize that the evolved

strain may have higher efficiency of RuBisCO by maintaining high intracellular CO2 concentration. During salinity stress, the redox status of an organism is disturbed (Shetty et al., 2019), leading to production of reactive oxygen (ROS) and nitrogen (RNS) species, which can degrade proteins and be harmful to the organism (Airaki et al., 2012; Bai et al., 2011). The Ev strain exhibited upregulation of nitric oxide reductase (NOR, cyp55b1) protein, but down-regulation of ROS scavenging enzymes ascorbate peroxidase, catalase and glutathione-S-transferase (Table 3.3). In C. reinhardtii, NOR is involved in NO scavenging and has been shown to reduce NO to nitrous oxide under hypoxic condition (Chang et al., 2011; Shoun, Fushinobu, Jiang, Kim, & Wakagi, 2012)). Furthermore, a NOR mutant strain of C. reinhardtii reduced nitrous oxide production by 70-90% (Plouviez et al., 2017). Interestingly, a recent study showed that nitric oxide signaling played an essential role in salinity response of C. reinhardtii (X. Chen et al., 2016). Increased NO production in C. reinhardtii during light stress has also been linked to autophagy and cell death under high light stress (Kuo, Chang, Lin, & Lee, 2020). As a result, it may be important for C. reinhardtii to upregulate NO scavenging mechanisms under stress conditions to avoid cell death. In our proteome analysis, NOR was upregulated by several fold (~5 fold) after long-term salinity acclimation, making NOR the highest upregulated protein in the evolved strain. On the other hand, ROS production does not seem to be an issue in the Ev strain, since several of the ROS detoxification enzymes were downregulated. Salinity increases osmotic stress in photosynthetic organisms, and many salinity tolerant strains, such as Dunaliella salina and C. sp. UWO241, upregulate the compatible solute glycerol production to alleviate osmotic stress (Goyal, 2007a; Kalra et al., 2020; Raymond, Morgan-Kiss, & Stahl-Rommel, 2020). Starch breakdown has been shown to support enhanced glycerol synthesis in different photosynthetic organisms (Goyal, 2007b). Several proteins involved in starch breakdown (Alpha-amylase, starch phosphorylase A and B) and starch synthesis (plastidal starch synthase, phosphoglucomutase, FBPase) were upregulated in both Ev-HS and Wt-HS. Furthermore, glycerol-3-phosphate dehydrogenase (GPDH), the enzyme responsible for conversion of dihydroxyacetone phosphate to glycerol-3-phosphate in the glycerol synthesis

122 pathway, was also highly upregulated under both the conditions. This transient starch metabolism in C. reinhardtii may help feed into glycerol biosynthesis during salinity acclimation, regulating osmotic stress. Contradictory to this result, the study by Perrineau et al. found upregulation of GPDH genes only in the parent strain after short-term salinity acclimation but not in the mixotrophically grown salinity evolved strain of C. reinhardtii (Perrineau et al., 2014). Surprisingly, the fermentation protein aldehyde/alcohol dehydrogenase (ADH1) was also upregulated during both short- and long-term salinity acclimation. ADH1 is a dual functioning enzyme that participates in the fermentative reduction of acetaldehyde or acetyl-CoA into ethanol and is crucial under anoxic conditions (Magneschi et al., 2012). However, it was recently shown that ADH1 can accumulate under oxic conditions, and increased levels of ADH1 coincide with increased starch production (Van Lis et al., 2017). Although ADH1 has been shown to play a role in anaerobic fermentation, it is not fully understood whether ADH1 also plays a role under oxic conditions. ADH1 levels were also shown to be upregulated under other abiotic stress conditions such as nitrogen and zinc deficiency (Van Lis et al., 2017). Thus, ADH1 may play a regulatory role in abiotic stress response. Moreover, a recent study showed that ADH1 confers salinity resistance in the cyanobacterial Synechocystis species (Yi et al., 2017). As starch is shown to be concomitantly upregulated with ADH1 levels (Van Lis et al., 2017), we suggest that dynamic regulation of starch synthesis/breakdown may help feed into the ADH1 mediated ethanol biosynthetic pathway conferring salinity resistance in the evolved strain. Starch biosynthesis and degradation pathways that are upregulated in the evolved strain after long-term salinity acclimation require not only fixed carbon but also increased ATP availability. From our photobiology data, we found that the evolved strain not only displays significantly higher oxygen evolution and CEF rates, but also shows high respiration rates compared to the parent strain. Both CEF and the respiration pathway provide extra ATP needed for starch metabolism. To this end, we found evidence for increased respiration in our proteome: several enzymes involved in the TCA cycle (MDH, ACH, IDH) as well as oxidative phosphorylation (ASA1, ASA3 and COX2B) were upregulated in the evolved strain. As a result, we suggest the evolved strain may use increased ATP generated from respiration and CEF to feed into starch metabolism. Photosynthetic organisms experience a variety of stresses in their natural habitat, which can be either short-term or long-term. Although much research has been done on short-term

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stresses, there are few studies done on the role of photosynthesis in long-term acclimation response. C. reinhardtii is a well-studied model organism for photosynthetic research with a sequenced genome and availability of many genetic tools (Merchant et al., 2007); however, its tolerance to many abiotic stress is fairly minimal. To this end, we believe C. reinhardtii is an ideal candidate to study stress adaptation response in the laboratory. Although C. reinhardtii is routinely grown in TAP media in the laboratory, the presence of acetate helps the organism to bypass photosynthetic carbon fixation during long-term stress acclimation (Perrineau et al., 2014). Under such conditions, it is hard to dissect the role that photosynthesis plays in adaptation when acetate is available. Furthermore, in the natural environment, green algae don’t always have access to readily available carbon to use during stress adaptation. Chlamydomonas algae are found in a myriad of environments, from freshwater and soil to snow and ice-covered lakes of polar habitats (Dolhi et al., 2013; Shetty et al., 2019). To survive in these different and sometimes extreme conditions, Chlamydomonas species need to restructure their photosynthetic and metabolic machinery to best suit their environment. Our study elucidates that green algae adapt to salinity stress by constitutively upregulating CEF, possessing high capacity for NPQ, and altering the downstream metabolism towards breakdown of starch to feed into glycerol and ethanol biosynthesis. Moreover, the salinity tolerant evolved strain downregulates wasteful mitochondrial photorespiration and may use CEF associated light-regulated glycolate oxidation instead. Lastly, we suggest that the

evolved strain maintains a high CO2 environment in the chloroplast, indicated by low levels of CCM proteins and upregulation of hypoxia-associated nitric oxide reductase protein. This response is quite different from the one observed under mixotrophic conditions, where the organism can bypass photosynthesis and use acetate as a source of energy (Perrineau et al., 2014). Further studies are needed to understand the exact relationship between CEF, photorespiration and CCM during salinity adaptation. The role of ADH1 mediated ethanol production in the evolved strain should also be explored to advance our understanding of metabolic changes associated with salinity adaptation. Last, future studies should focus on teasing apart the role of ROS vs RNS scavenging mechanisms in conferring resistance to salinity in C. reinhardtii.

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2.0 Wt-LS Wt-HS 1.5 Ev-LS

1.0 Ev-HS

O.D. (750 nm) 0.5

0.0 0 1 2 3 4 5 6 7 8 9 10 Time (days)

Figure 3.1. Comparison of growth of different C. reinhardtii strains. Wildtype (Wt) UTEX 89 was evolved under constant salinity pressure of 100 mM in BBM until an evolved strain (Ev) with growth rate comparable to the original wild type strain was achieved (48 transfers). Growth curve was conducted with Wt and Ev strain grown under both low salinity (LS, BBM) and high salinity (HS, BBM+100 mM NaCl) conditions. Optical density was measured at 750 nm. Data are representative of three independent samples and the standard error are shown (n=3)

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Wt-LS 0.8 A 1.0 B Wt-HS 0.6 Ev-LS Ev-HS 0.4 0.5 qL YPSII

0.2

0.0 0.0 0 200 400 600 800 0 200 400 600 800 1000 PAR (µE) PAR (µE)

1.0 C 4 D 0.8 3 0.6 2 NPQ

Y(NPQ) 0.4 1 0.2

0.0 0 0 200 400 600 800 0 200 400 600 800 PAR (µE) PAR (µE)

0.9 Wt-HS NO F 0.6 E 0.8 Wt-HS NPQ 0.5 0.7 Ev-HS NO 0.6 Ev-HS NPQ 0.4 0.5 0.3 0.4

Y(NO) Wt-LS 0.2 Wt-HS 0.3 Ev-LS 0.2 0.1 Yield (YNO/YNPQ) Ev-HS 0.1 0.0 0.0 0 200 400 600 800 0 200 400 600 800 PAR (µE) PAR (µE)

Figure 3.2. PSII parameters for parent (Wt) and evolved (Ev) strain under low (LS) and high (HS) salinity during a light curve. Photosystem II yield (YPSII, A), proportion of open PSII (qL, B), capacity for non- photochemical quenching (YNPQ, C), non-photochemical quenching (NPQ, D), capacity for non-regulated photochemical quenching (YNO, E), changes in non-regulated and regulated non-photochemical quenching (F). (mean ± SD, n =3)

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A B 12 ** 3 -1 10 -1 cell 8 cell 2 -1 -1

6 min min 2 2 4 1 nmol O nmol O 2

0 0 Wt-LS Ev-HS Wt-LS Ev-HS

Figure 3.3. Oxygen evolution (A) and dark respiration (B) rates of the parent strain in low salt (Wt-LS) and evolved strain (Ev-HS) in high salinity (100 mM NaCl). Data are representative of three independent samples and the standard error are shown (n=3)

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A 0.5 **** B 1250 **** *** *** 0.4 1000 *** **** 0.3 ** 750 820 (ms) A Δ

0.2 1/2 500 t

0.1 250

0.0 0

Wt-LS Wt-HS Ev-LS Ev-HS Wt-LS Wt-HS Ev-LS Ev-HS

Figure 3.4. P700 kinetics of parent and the evolved strain under low (LS) and high (HS) salinity. A. Proportion of photooxidizable P700, calculated as change in absorption of P700 at 820 nm under FR light. B. P700+ re-reduction time for all four conditions. Rate of re-reduction is calculated using exponential decay kinetics after FR is switched off. (**, p < 0.05; ***, p < 0.01; ****, p < 0.005; n =9).

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Ev-HS vs Wt-LS WT-HS vs WT-LS cyt P450/nitric oxide NAD(P)-bd_dom domain-containing protein NAD(P)-bd_dom domain-containing protein Alcohol Dehyrogenase DJ-1_PfpI domain-containing protein Zygote-specific Zys3 like protein FAD-binding protein Uncharacterized protein Protein disulfide isomerase Glycerol-3-phosphate dehydrogenase [NAD(+)] Nudix hydrolase 12 Alpha-amylase Alpha-1,4 glucan phosphorylase Glycerol-3-phosphate dehydrogenase [NAD(+)] 12 Alpha-amylase NAD(P) transhydrogenase Alcohol Dehydrogenase Isocitrate dehydrogenase DJ-1_PfpI domain-containing protein Glycerol-3-phosphate dehydrogenase [NAD(+)] Glycerol-3-phosphate dehydrogenase [NAD(+ UDP-Glucose:protein transglucosylase Uncharacterized protein 6-phosphogluconate dehydrogenase Ribosomal protein L24 Superoxide dismutase ATP-sulfurylase Alpha-1,4 glucan phosphorylase Glycerol-3-phosphate dehydrogenase [NAD(+)] Protein disulfide isomerase UDP-D-glucuronic acid decarboxylase 10 6-phosphogluconate dehydrogenase 10 Nascent polypeptide-associated complex subunit beta Isocitrate dehydrogenase [NADP] 40S ribosomal protein S7 Superoxide dismutase Nudix hydrolase Starch synthase Predicted protein S-adenosylmethionine synthase Uncharacterized protein Peptidylprolyl isomerase Uncharacterized protein Elongation factor EF-3 Aconitate hydratase UDP-Glucose:protein transglucosylase Glycerol-3-phosphate dehydrogenase [NAD(+) Alpha-1,4 glucan phosphorylase Cell wall protein pherophorin-C2 Malate dehydrogenas Cell wall protein pherophorin-C13 8 Isocitrate dehydrogenase [NAD] subunit 8 Tautomerase domain-containing protein UDP-D-glucuronic acid decarboxylase Ribosomal protein L3 Lhcb3 HP domain-containing protein Malate dehydrogenase Glutathione-S-transferase Ribosomal protein S19 S-adenosylmethionine synthase Heat shock protein 90A Predicted protein Vasa intronic gene Calreticulin Histone H2A Cell wall glycoprotein GP2 (Fragment) Rhodanese-like protein Histone H2A 6 Uncharacterized protein Histone H2A Starch synthase, chloroplastic/amyloplastic 6 Histone H2A Elongation factor T Histone H2A Lhcb5 Fructose-1,6-bisphosphatase FAS1 domain-containing protein CAAD domain-containing protein Cytochrome c oxidase subunit II Ribosomal protein L19 HP domain-containing protein Lhcb5 Alpha-1,4 glucan phosphorylase Lhcb3 40S ribosomal protein S6 Ubiquinol:cytochrome c oxidoreductase LhcI-2 ATP syntase-associated protein ASA1 4 Porphobilinogen deaminase Fructose-1,6-bisphosphatase Oxygen-evolving enhancer protein 1 4 Mitochondrial chaperonin 60 Protein disulfide-isomerase Uncharacterized protein Oxygen evolving enhancer protein 3 40S ribosomal protein S4 ATP synthase associated 36.3 kDa protein Uncharacterized protein Glycine cleavage system P protein NADPH-protochlorophyllide oxidoreductase Enolase Fructose-bisphosphate aldolase Nop domain-containing protein PsaD 4-hydroxy-tetrahydrodipicolinate synthase Lhca Phosphoglucomutase FBPase domain-containing protein 2 Pyruvate kinase 2 Plastid ribosomal protein L19 Histone H4 PKS_AT domain-containing protein Biotin carboxylase PETC Serine hydroxymethyltransferase Rubisco activase Elongation factor G 30S ribosomal protein S3 Nitrogen regulatory protein PII Uncharacterized protein PSII_BNR Histone H4 Serine hydroxymethyltransferase CoA carboxyltransferase CoA carboxyltransferase Vacuolar proton pump Glycine cleavage system P Dihydrolipoamide acetyltransferase of PDH complex 0 Nucleoside diphosphate kinase 0 30S ribosomal protein S3 Cytochrome f RuBisCO small Rubisco activase RuBisCO small 4-hydroxy-tetrahydrodipicolinate synthase Mg protoporphyrin IX trransferase ribosomal protein L4 Ribosomal protein L23a Biotin carboxylase Peptidylprolyl isomerase ribosomal protein L3 ADP ribosylation factor-like 3 NAC-A/B domain-containing protein 3-oxoacyl-[acyl-carrier-protein] synthase ATP-grasp domain-containing protein Chaperonin 11 Peptidyl-prolyl cis-trans isomerase -2 -2 PsaC Uncharacterized protein Glycine--tRNA ligase Uncharacterized protein Phosphoglycerate kinase Uncharacterized protein Acetohydroxyacid dehydratase Inosine-5'-monophosphate dehydrogenase Hydroxypyruvate reductase Thioredoxin domain-containing protein GrpE protein homolog Phosphoribosylamine--glycine ligase (Fragment) Uncharacterized protein Nucleoside diphosphate kinase Low-CO2 inducible protein Spermine synthase Uncharacterized protein Pet O subunit cytb6f Hydroxypyruvate reductase Chaperonin 11 -4 -4 Nucleic acid binding protein Glutathione S-transferase Chlamyopsin 2 Nop domain-containing protein L-ascorbate peroxidase GrpE protein homolog Catalase Histone H4 GrpE protein homolog Ribose-5-phosphate isomerase ATP-grasp domain-containing protein Thylakoid lumenal 17.4 kDa protein Nucleic acid binding protein GrpE protein homolog PetC Uncharacterized protein Uncharacterized protein Peptidylprolyl isomerase Phototropin -6 Adenylylphosphosulfate reductase -6 Uncharacterized protein Uncharacterized protein Predicted protein SPS1p Elongation factor P Glyoxylate/succinic semialdehyde reductase Low-CO2 inducible protein Adenylylphosphosulfate reductase 38 kDa RNA-binding protein Uncharacterized protein FAD-binding FR-type domain-containing protein Plastid ribosomal protein S6 Phosphoribosylamine--glycine ligase 3-oxoacyl-reductase FAD-binding FR-type domain-containing H/ACA ribonucleoprotein complex subunit Glycolate dehydrogenase LL-diaminopimelate aminotransferase Ribosomal protein S29 -8 Uncharacterized protein -8 Glutathione S-transferase Predicted protein Peptidylprolyl isomerase LCIB_C_CA domain-containing protein Glycolate dehydrogenase PeTO LCIB-like gene Flagellar associated protein Uncharacterized protein TPM_phosphatase domain-containing protein Heme oxygenase 50S ribosomal protein L23 LCiD Low-CO2 inducible protein Predicted protein -10 TPM_phosphatase domain-containing protein -10 FC 1 FC 2 FC 3 FC 1 FC 2 FC 3

Figure 3.5. Heat map of proteins that are significantly affected under acclimation (WT-HS) and evolution (Ev-HS) under high salinity compared to the parent strain (WT-LS). Protein identities are shown on left and each column indicates the fold change (FC) difference between the test and the parent strain (p < 0.05) for each biological replicate (n =3). Proteins that are upregulated are in yellow-orange spectrum and proteins that are downregulated are in red-purple spectrum.

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Figure 3.6. Analysis of significantly differentially regulated proteins of Wildtype (Wt) and Evolved (Ev) strain under high salinity (HS) compared to Wildtype strain under low salinity. A. Venn diagram of significantly differentially regulated proteins (Tfold, Pattenlab 4.0) that are common and unique (A) in Ev-HS and Wt-HS. B. The cellular localization pattern of those proteins in the two strains. The up-regulated (pink) and the down-regulated (blue) proteins of both Wt-HS (dark color) and Ev-HS (light color) are shown. The total number of proteins belonging to each cellular location are shown as a sum of up- and down-regulated proteins on the x-axis. (n=3, p<0.05)

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GTP binding Oxygen evolving activity Wt-HS Up Transaminase activity Wt-HS Down Elongation factor Chaperone binding Ev-HS Up Kinase Ev-HS Down Electron transfer activity Ligase Unfolded protein binding Carboxylase Metal ion binding Actin binding Transcription factor binding DNA binding Chlorophyll binding Lyase ATPase activity ATP binding Ribosomal protein NAD binding Transferase RNA binding Glycogen Phosporylase Hydrolase Isomerase Dehydrogenase activity Oxidoreductase Dehydratase 0 5 10 15 20 25 Significantly differentially expressed proteins

Figure 3.7. Categorization of significantly differentially regulated proteins (Tfold, Patternlab 4.0) of wildtype strain (Wt-HS) and evolved strain (Ev-HS) under high salinity into Gene Ontology (GO) molecular function. Proteins belonging to each sub-category are differentiated into up-regulated (pink) and down-regulated (blue) for both Wt-HS (dark color) and Ev-HS (light color). Total number of proteins associated with each sub-category are shown as a sum of up- and down-regulated proteins on the x-axes (n=3, p < 0.05).

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Starch and sucrose metabolism Ev-HS Up Sugar metabolism Ev-HS Down Lipid metabolism Fermentation Wt-HS Up Nitric oxide Signalling Wt-HS Down Oxidative Stress Chaperones and folding catalyts Pentose Phosphate pathway Glutathione metabolism TCA cycle Sulfur metabolism Carbon fixation Photosynthesis Cellular processes and transport Cysteine and methionine metabolism Secondary metabolism Protein biosynthesis Glycolysis/Gluconeogenesis Porphyrin and chlorophyll metabolism Pyruvate metabolism Chromosome and associated proteins Fatty acid biosynthesis Glyoxylate and dicarboxylate metabolism Nucleotide biosynthesis Lysine biosynthesis Carbon concentrating mechanism Phototaxis/Motility One-carbon metabolism 0 5 10 15 20 Significantly differentially expressed proteins

Figure 3.8. Categorization of significantly differentially regulated proteins of Wildtype (Wt-HS) and Evolved (Ev-HS) under high salinity into Kegg Orthology (KO) biological pathway. Proteins belonging to each sub-category are differentiated into up-regulated (pink) and down-regulated (blue) for both Wt-HS (dark color) and Ev-HS (light color). Total number of proteins associated with each sub-category are shown as a sum of up- and down-regulated proteins on the x-axes (n=3, p < 0.05).

132

A Up-regulated 10

8 Wt-HS Ev-HS 6

2 Fold Change (HS vs LS) 0

Alpha amylase Nudix hydrolase Cytochrome P450

Superoxide dismutase Malate dehydrogenase Uncharacterized protein Isocitrate dehydrogenase Protein disulfide isomeraseSulfate adenylyltransferase UDP-arabinopyranose mutase HP domain-containing protein StarchD-Fructose-1,6-bisphosphatase synthase,Chlorophyll chloroplasticChlorophyll a-b binding a-b binding protein protein Alpha-1,4-glucan phosphorylase Alpha-1,4-glucan phosphorylase S-adenosylmethionine synthase Aldehyde-Alcohol dehydrogenase DJ-1_PfpI domain-containing protein 6-phosphogluconate dehydrogenase Glycerol-3-phosphateGlycerol-3-phosphate DehydrogenaseGlycerol-3-phosphate Dehydrogenase Dehydrogenase UDP-D-glucuronic acid decarboxylase NAD-dependent epimerase/dehydratase Tautomerase domain-containing protein Down-regulated 0

-2

-4

-6

-8 Wt-HS Ev-HS Fold Change (HS vs LS) -10

Chaperonin 11 Chlamyopsin 2 Rubisco Activase Heme Oxygenase Predicted protein Predicted protein Biotin carboxylase Spermine Synthase GrpE protein homolog GrpE protein homolog UncharacterizedUncharacterized protein protein CoA carboxyltransferase Peptidylprolyl isomerase Glutathione S-transferase Glycolate dehydrogenase Nitrogen regulatoryHydroxypyruvate protein reductase Nucleic acid binding protein Nucleoside diphosphate kinase Low CO2 inducibleNop protein domain-containing LCIC protein Low CO2 inducibleLow protein CO2 inducibleLCIE protein LCID Serine hydroxymethyltransferase Glycine cleavage system P protein AdenylylphosphosulfateFormyltetrahydrofolate reductase synthetaseCytochrome b6f complex subunit V

30S ribosomal protein S3, chloroplastic Diadenosine tetraphosphate synthetase Glycinamide ribonucleotide synthetase 4-hydroxy-tetrahydrodipicolinate synthase PETC subunit of the cytochrome b6f complex TPM_phosphatase domain-containing protein FAD-binding FR-type domain-containing protein

Figure 3.9. Analysis of set of common proteins that are significantly differentially regulated in Wildtype (Wt-HS) and Evolved strain (Ev-HS) under high salinity compared to the parent strain in low salinity. Wt-HS (black) and Ev-HS (gray) share 64 common proteins that are upregulated (A) and downregulated (B) out of 100 and 124 proteins total. Fold change of Wt-HS or Ev-HS as compared to Wt-LS is shown on the y-axis (n=3, p<0.05). The uniport identities of each identifies protein is shown on the x-axis.

133

Evolved-HS A 8 6

4 2 0

Fold change -2 -4 -6

Lhca PsaC Enolase Catalase PSII_BNR Histone H2AHistoneHistone H2AHistone H2AHistone H2A H2A RuBisCORuBisCO small small NAC subunit beta Vasa intronic gene Aconitate hydratase Elongation factor G Elongation factor P Ribosomal protein L3 ribosomal protein L4 3-oxoacyl-reductase Ribosomal proteinElongation L24 factor EF-3 Vacuolar proton pump RibosomalHeat shock protein protein S19 90A L-ascorbate peroxidaseUncharacterized protein Peptidylprolyl isomerase 40S ribosomal protein40S ribosomal S6 protein S4 Phosphoglycerate kinase NAD(P) transhydrogenase40S ribosomal protein S7 50S ribosomal protein L23 Plastid ribosomal protein L3 Mitochondrial chaperoninPlastid 60 ribosomal protein L19 38 kDaPlastid RNA-binding ribosomal protein protein S6 Acetohydroxyacid dehydratase Cell wall protein pherophorin-C2 Ribose-5-phosphate isomerase Cell wall protein pherophorin-C13 FBPase domain-containing protein Mg protoporphyrin IX trransferase PKS_AT domain-containing protein NAC-A/B domain-containing protein ThylakoidPeptidylprolyl lumenal 17.4 isomerase kDa protein FKBP type Cell wall glycoprotein GP2 (Fragment)ATP syntase-associated protein ASA1 LL-diaminopimelate aminotransferase Ubiquinol:cytochrome c oxidoreductase Thioredoxin domain-containing protein ATP synthase associated 36.3 kDa protein H/ACA ribonucleoprotein complex subunit

Dihydrolipoamide acetyltransferasePeptidyl-prolyl of PDH complexcis-trans isomerase (cyclophilin type)

134

B 8 Wildtype-HS

6 4 2 0

Fold change -2 -4 -6

Calreticulin Histone H4Histone H4 Histone H4Phototropin

Predicted protein Pyruvate kinase Predicted protein Pyruvate carboxylase Ribosomal protein L19 Phosphoglucomutase Ribosomal protein S29 UncharacterizedUncharacterized protein Rhodanese-like proteinUncharacterized protein protein UncharacterizedRibosomal protein protein L23a Uncharacterized protein Glutathione-S-transferase Protein disulfide-isomerase Flagellar associated protein ADP ribosylation factor-like 3 Hydroxymethylbilane synthase Zygote-specific Zys3 like protein CAAD domain-containing protein Fructose-bisphosphate aldolase Isocitrate dehydrogenase [NADP]Elongation factorLight Ts, harvesting mitochondrial complex I (Lhca3) Oxygen-evolvingOxygen enhancer evolving proteinenhancer 1 protein 3 3-oxoacyl-[acyl-carrier-protein] synthase NADPH-protochlorophyllide oxidoreductase Inosine-5'-monophosphate dehydrogenase Photosystem I reaction center subunit II (PsaD)

Figure 3.10. Analysis of set of unique proteins that are significantly differentially regulated in Wildtype (Wt-HS) and Evolved strain (Ev-HS) under high salinity compared to the parent strain in low salinity. The evolved strain has a set of 60 proteins that are uniquely differentially regulated under high salinity (A) whereas the Wt-HS differentially regulates only 24 unique proteins (B). Fold change of Wt-HS or Ev-HS as compared to Wt-LS is shown on the y-axis (n=3, p<0.05). The uniport identities of each identifies protein is shown on the x- axis.

135

Table 3.1. Growth rates, doubling time and final biomass of C. reinhardtii wildtype (Wt) and evolved (Ev) strains under low (LS) and high (HS) salinity. Data are representative of three independent samples and the standard error are shown (n=3; average ± SD)

Strain Growth rate (hr-1) Doubling Time Final Biomass

(hr) (O.D.750) Wt-LS 0.0088±0.0008 68.87±4.73 1.33±0.233

Wt-HS 0.0057±0.0010 81.29±9.31 0.69±0.153

Ev-LS 0.0104±0.0004 53.43±3.99 1.86±0.157

Ev-HS 0.0083±0.0004 69.79±1.34 1.2±0.106

136

Table 3.2. Significantly differentially regulated proteins in Chlamydomonas reinhardtii wildtype under high salinity (Wt-HS) compared to parent strain under low salinity (Wt-LS). Identified proteins are divided into KEGG functional categories and organelle localization is provided based on Uniprot prediction. Fold change and p-value are shown (Tfold, Patternlab). Differentially regulated proteins that are unique to the strain are signified with an asterisk(*).

Protein name Uniprot KB Fold Change p-Value Sub-cellular Gene (vs. Wt-LS) localization Photosynthesis Chlorophyll a-b Q9FEK6 1.64820628 0.00548312 Chloroplast Lhcb5 binding protein Chlorophyll a-b Q9ZSJ4 1.59618773 0.00283989 Chloroplast Lhcb3 binding protein Chlorophyll a-b Q75VY9 1.53509345 0.02032394 Chloroplast Lhca3 binding protein* Oxygen-evolving A8J0E4 1.4902581 0.01303625 Chloroplast PsbO enhancer protein 1 of photosystem II* Oxygen evolving A8JEV1 1.42375615 0.01463701 Chloroplast PsbQ enhancer protein 3* Photosystem I Q5NKW4 1.33905571 0.00867464 Chloroplast PsaD reaction center subunit II, 20 kDa* Cytochrome f P23577 -1.4864801 0.00912624 Chloroplast PetA Rieske iron-sulfur A8J9Y1 -2.0125831 0.01023433 Chloroplast PetC subunit of the cytb6f Cytochrome b6f A8JGW2 -3.0817928 0.00427499 Chloroplast PetO complex subunit V Carbon fixation

137

D-fructose-1,6- A8IKQ0 1.76167819 0.00413978 Cytosol; FBP1 bisphosphate 1- chloroplast phosphohydrolase Fructose- A8JCY4 1.38664111 0.01078353 Cytosol FBA1 bisphosphate aldolase* Rubisco activase Q6SA05 -1.5130153 0.00196201 Rca Carbon concentrating mechanism Peptidylprolyl A0A2K3D521 -1.6023369 0.00510904 Chloroplast FKB19 isomerase Low-CO2 inducible Q75NZ1 -1.8032067 0.02989841 LciC protein LCIB_C_CA A0A2K3DUD -3.0742711 0.00759625 LCIE domain-containing 5 protein Pentose phosphate pathway 6-phosphogluconate A8J5F7 3.18609768 8.39E-05 Cytosol GND1a/G dehydrogenase ND1b D-fructose-1,6- A8IKQ0 1.76167819 0.00413978 Cytosol; FBP1 bisphosphate 1- chloroplast phosphohydrolase Fructose- A8JCY4 1.38664111 0.01078353 Cytosol FBA1 bisphosphate aldolase* Phosphoglucomutase A8J8Z1 1.3350703 0.00503698 Cytosol GPM1a|G (alpha-D-glucose-1,6- PM1b bisphosphate- dependent) Glycolysis/Gluconeogensis

138

D-fructose-1,6- A8IKQ0 1.76167819 0.00413978 Cytosol; FBP1 bisphosphate 1- chloroplast phosphohydrolase Fructose- A8JCY4 1.38664111 0.01078353 Cytosol FBA1 bisphosphate aldolase* Phosphoglucomutase A8J8Z1 1.3350703 0.00503698 Cytosol GPM1a|G (alpha-D-glucose-1,6- PM1b bisphosphate- dependent) Pyruvate kinase A8IVR6 -1.3376174 0.00828916 PYK1 Pyruvate metabolism Pyruvate kinase A8IVR6 -1.3376174 0.00828916 PYK1 Pyruvate A0A2K3DMK -1.8251207 0.01791654 Cytosol PYC1 carboxylase* 8 Glycolate A8J2E9 -2.6622485 0.00629615 GYD1 dehydrogenase TCA cycle Isocitrate A0A2K3DTT8 3.08570607 0.01189051 Mitochondrion IDH3 dehydrogenase [NADP]* Malate P93106 2.39251936 0.00182673 Mitochondrion MDH2 dehydrogenase Isocitrate A0A2K3E3Z0 2.39242833 0.00057603 Mitochondrion IDH2 dehydrogenase [NAD] Pyruvate A0A2K3DMK -1.8251207 0.01791654 Cytosol PYC1 carboxylase* 8 Glutathione metabolism

139

6-phosphogluconate A8J5F7 3.18609768 8.39E-05 Cytosol GND1a/G dehydrogenase ND1b Isocitrate A0A2K3DTT8 3.08570607 0.01189051 mitochondrion IDH3 dehydrogenase [NADP]* Glutathione-S- A8JBA7 2.20378638 0.0265636 GSTS1 transferase* Glutathione-S- A8JBB4 -1.8364959 0.02303077 GSTS2 transferase Starch and sucrose metabolism Alpha-amylase A0A2K3DGK 4.01347971 0.00102383 AMYA2 5 Alpha-1,4-glucan A8IYK1 3.37556578 0.00067464 PHOB phosphorylase Alpha-1,4-glucan Q2VA40 2.56893091 0.00030462 PHOA phosphorylase Starch synthase, A0A2K3CQC 1.80517565 0.02623777 Chloroplast STA2 chloroplastic 6 Phosphoglucomutase A8J8Z1 1.3350703 0.00503698 Cytosol GMP1a/1 b Lipid metabolism Glycerol-3-phosphate A0A2K3E871 4.06243764 0.00155464 Cytosol CHLRE_0 dehydrogenase 1g053150 [NAD(+)] v5 Glycerol-3-phosphate A0A2K3E866 3.95034909 0.00115418 Cytosol CHLRE_0 dehydrogenase 1g053150 [NAD(+)] v5 Glycerol-3-phosphate A0A0B5KTL4 3.54368459 0.00077889 Cytosol CHLRE_0 dehydrogenase 1g053150 [NAD(+)] v5

140

Fatty acid biosynthesis Biotin carboxylase A0A2K3DGE -1.3826233 0.00113906 CHLRE_0 7 8g359350 v5 CoA A0A2K3D1L8 -1.4216686 0.01819546 CHLRE_1 carboxyltransferase 2g484000 N-terminal domain- v5 containing protein 3-oxoacyl-[acyl- A0A2K3D7U3 -1.6635588 0.01238204 CHLRE_1 carrier-protein] 1g467723 synthase v5 Fermentation Aldehyde-alcohol A8JI07 3.86061197 0.00453928 Mitochondrion ADH1 dehydrogenase Sugar metabolism NAD-dependent A0A2K3E0N9 7.94625197 1.00E-05 SNE5 epimerase/dehydratas e UDP- A2PZC2 2.67664613 0.00193355 Cytosol UPTG1 arabinopyranose mutase UDP-D-glucuronic A8IEW6 2.35140038 0.00252446 GAD1 acid decarboxylase Phosphoglucomutase A8J8Z1 1.3350703 0.00503698 Cytosol GMP1a/1 b Sulfur metabolism ATP-Sulfurylase A0A2K3DZ59 2.8081399 0.00091595 ATS1 Adenylylphosphosulf A8J6A7 -2.1949001 0.00475618 MET16 ate reductase Nitric oxide signaling

141

Cytochrome P450, A0A2K3E587 3.46336222 0.00338099 CYP55B1 nitric oxide reductase Nitrogen regulatory A0A2K3DLS9 -1.7726453 0.00978464 Cytosol GLB1 protein PII Oxidative Stress Superoxide dismutase A8JAX1 3.39883461 0.00667099 Mitochondrion MSD2 Chaperones and folding catalysts Protein disulfide A8HQT1 3.22621103 0.0054228 ER PDI2 isomerase Calreticulin* A8HMC0 2.02882976 0.02220176 ER CRT2 Uncharacterized A0A2K3DG79 1.90110197 0.02899693 Chloroplast; protein mitochondrion Protein disulfide- O48949 1.42647286 0.00525114 ER PDI isomerase Chaperonin 11 A8J3C3 -1.6651462 0.02916665 Chloroplast; CPN11 mitochondrion GrpE protein Q945T2 -1.7447274 0.00128448 CGE1 homolog GrpE protein Q945T1 -1.86875 0.00031511 CGE1 homolog Amino acid metabolism S- A0A2K3DLX 2.11989935 0.01687047 Cytosol METM adenosylmethionine 7 synthase Uncharacterized A0A2K3DKV 1.42134316 0.02075588 CHLRE_0 protein 5 7g344400 v5 4-hydroxy- A0A2K3E2N0 -1.5392797 0.00297714 DPS1 tetrahydrodipicolinate synthase

142

Protein Biosynthesis Elongation factor Ts* A0A2K3D3Y7 1.79733008 0.00415134 Mitochondrion EFTS

Ribosomal protein A8IA18 1.68135545 0.01357651 Cytosol RPL19 L19 30S ribosomal protein Q08365 -1.3988092 0.01415078 Chloroplast RPS3 S3 Uncharacterized A0A2K3E7Z9 -1.4019389 0.00497687 CHLRE_0 protein* 1g050316 v5 Ribosomal protein A8J239 -1.5732339 0.01270483 Cytosol RPL23a L23a Uncharacterized A0A2K3DRT -1.7067233 0.00403867 Mitochondrion CHLRE_0 protein 1 5g233800 v5 NOP domain A0A2K3D4E7 -1.8586356 0.00031417 Nucleus NOP56 containing protein Ribosomal protein A8JIE5 -2.6642573 0.01037559 Chloroplast RPS29 S29* Cellular processes and transport HP domain- A0A2K3D4C8 2.33012814 0.00652927 CHLRE_1 containing protein 2g524400 v5 Chlamyopsin A0A2K3E4T1 -1.7338575 0.00646438 cop 2/Babo1

143

Table 3.3. Significantly differentially regulated proteins in Chlamydomonas reinhardtii evolved strain under high salinity (Ev-HS) compared to parent strain under low salinity (Wt-LS). Identified proteins are divided into KEGG functional categories and organelle localization is provided based on Uniprot prediction. Fold change and p-value are shown (Tfold, Patternlab). Differentially regulated proteins that are unique to the strain are signified with an asterisk(*).

Protein name Uniprot Fold Change p-Value Sub-cellular Gene

KB (vs. Wt-LS) localization

Photosynthesis Chlorophyll a-b Q9FEK6 1.702329656 0.011703666 Chloroplast Lhcb5 binding protein Chlorophyll a-b Q9ZSJ4 1.843519394 0.015935159 Chloroplast Lhcb3 binding protein Chlorophyll a-b Q84Y02 -1.404415719 0.01630453 Chloroplast Lhca binding protein Rieske iron-sulfur A8J9Y1 -1.523441643 0.021377076 Chloroplast PetC subunit of the cytb6f Photosystem I core Q00914 -1.713110762 0.014538899 Chloroplast PsaC protein* Cytochrome b6-f A8JGW2 -1.888423853 0.018923534 Chloroplast PetO complex subunit petO Carbon fixation D-fructose-1,6- A8IKQ0 1.513365284 0.012271092 Chloroplast, FBP1 bisphosphate 1- cytosol phosphohydrolase Sedoheptulose-1,7- A0A2K3D -1.48655124 0.00465111 Chloroplast SEBP1 bisphosphatase* Y10 Rubisco activase Q6SA05 -1.524854346 0.007761717 Chloroplast RCA RuBisCO small* A0A2K3E -1.64195739 0.012461088 Chloroplast RBCS2 3L3 RuBisCO small* P00873 -1.64195739 0.012461088 Chloroplast RBCS1 Phosphoglycerate A8JC04 -1.739404757 0.011436339 Cytosol PGK1 kinase*

144

Ribose-5-phosphate A8IRQ1 -2.054677376 0.017604485 RPI1 isomerase* Carbon concentrating mechanism Low-CO2 inducible Q75NZ1 -2.291702003 0.011363177 Chloroplast LCIC protein Peptidylprolyl A8IRU6 -1.700711529 0.013345218 CYN38 isomerase cyclophilin type Peptidylprolyl A0A2K3D -3.432043929 0.000707149 Chloroplast FKB19 isomerase 521 LCIB-like gene A0A2K3D -4.011249939 0.004596238 LCIE UD5 Low-CO2 inducible Q0Z9B8 -4.684664998 0.001745959 LCID protein Photorespiration Glycolate A8J2E9 -3.590740733 0.003822894 GYD1 dehydrogenase Pentose phosphate pathway 6-phosphogluconate A8J5F7 2.565092979 0.00020223 Cytoplasm GND1A, dehydrogenase GND1B D-fructose-1,6- A8IKQ0 1.513365284 0.012271092 Chloroplast FBP1 bisphosphate 1- phosphohydrolase Ribose-5-phosphate A8IRQ1 -2.054677376 0.017604485 RPI1 isomerase* Glycolysis/Gluconeogensis D-fructose-1,6- A8IKQ0 1.513365284 0.012271092 Chloroplast FBP1 bisphosphate 1- phosphohydrolase Pyruvate metabolism

145

Phosphopyruvate A8JH98 -1.380153706 0.007483646 PGH1 hydratase* Dihydrolipoamide A0A2K3D -1.597832876 0.02464924 DLA2 acetyltransferase of W88 PDH complex* Phosphoglycerate A8JC04 -1.739404757 0.011436339 Cytosol PGK1 kinase* TCA cycle Isocitrate A0A2K3E 2.704907782 0.005963003 Mitochondrion IDH2 dehydrogenase 3Z0 [NAD+] Aconitate hydratase* A0A2K3E 2.000972041 0.015958437 Mitochondrion ACH1 7I5 Malate dehydrogenase P93106 1.843195477 0.001398694 Mitochondrion MDH2 Dihydrolipoamide A0A2K3D -1.597832876 0.02464924 DLA2 acetyltransferase of W88 PDH complex* Oxidative phosphorylation Cytochrome c oxidase Q9AU02 1.62019464 0.019329778 Cox2B subunit II ATP synthase- A8ITL0 1.550223403 0.004964928 Mitochondrion ASA1 associated protein ASA1 ATP synthase A8JCE9 1.404396678 0.020945933 Mitochondrion ASA3 associated 36.3 kDa protein Glutathione metabolism 6-phosphogluconate A8J5F7 2.565092979 0.00020223 Cytosol GND1A, dehydrogenase GND1B

146

Glutathione S- A8JBB4 -3.259072563 0.004923126 GSTS2 transferase Starch and sucrose metabolism Alpha-amylase A0A2K3D 2.808377597 0.00047674 AMYA2 GK5 Alpha-1,4-glucan A8IYK1 2.890105405 0.0025806 PHOB phosphorylase Alpha-1,4-glucan Q2VA40 1.598382186 0.00768859 PHOA phosphorylase Starch synthase, A0A2K3C 2.252303853 0.00400426 Chloroplast STA2 chloroplastic/Amylopla QC6 stic Lipid metabolism Glycerol-3-phosphate A0A2K3E 2.68568117 0.00229253 Cytosol CHLREDR dehydrogenase 871 AFT_1469 [NAD(+)] 45 Glycerol-3-phosphate A0A2K3E 2.43896031 0.00370424 Cytosol CHLRE_0 dehydrogenase 866 1g053150v [NAD(+)] 5 Glycerol-3-phosphate A0A0B5K 1.97240287 0.00790768 Cytosol CHLRE_0 dehydrogenase TL4 1g053000v [NAD(+)] 5 Fatty acid biosynthesis FAD-binding FR-type A0A2K3E 3.278980259 0.001979404 ER CHLRE_0 domain-containing 4Y8 1g004900v protein* 5 PKS_AT domain- A0A2K3C -1.503008983 0.021194887 CHLRE_1 containing protein* Y26 4g621650v 5

147

CoA A0A2K3D -1.596170839 0.011116055 CHLRE_1 carboxyltransferase 1L8 2g484000v 5 Biotin carboxylase A0A2K3D -1.665454728 0.000294921 CHLRE_0 GE7 8g359350v 5 3-oxoacyl-reductase* Q84X75 -2.666501309 0.016577451 Chloroplast CHLRE_0 3g172000v 5 Sugar metabolism NAD-dependent A0A2K3E 4.00892672 0.00558223 SNE5 epimerase/dehydratase 0N9 UDP-arabinopyranose A2PZC2 2.610136482 0.00381745 UPTG1 mutase UDP-D-glucuronic A8IEW6 2.382794963 0.00272116 GAD1 acid decarboxylase Fermentation Aldehyde-alcohol A8JI07 3.550383218 0.000239034 Mitochondrion ADH1 dehydrogenase Sulfur metabolism ATP-sulfurylase A0A2K3D 2.541426136 0.01017567 ATS1 Z59 Adenylylphosphosulfat A8J6A7 -2.182869648 0.001830861 MET16 e reductase Nitric oxide signalling Cytochrome P450, A0A2K3E 4.971052197 0.000298635 CYP55B1 nitric oxide reductase 587 Nitrogen regulatory A0A2K3D -1.55159014 0.023318946 Cytosol GLB1 protein PII LS9 Oxidative Stress

148

Superoxide dismutase A8JAX1 2.29175518 0.012322674 Mitochondrion MSD2 L-ascorbate A8JG56 -1.948733871 0.021257698 CHLRE_0 peroxidase* 5g233900v 5 Catalase* A8J537 -1.953152615 0.018064525 CAT1 Glutathione S- A8JBB4 -3.259072563 0.004923126 GSTS2 transferase Chaperones and folding catalysts Protein disulfide A8HQT1 3.035370024 0.000426994 PDI2 isomerase Peptidylprolyl A8HUK0 2.104454655 0.0108878 FKB12 isomerase* Heat shock protein A8J1U1 1.783499204 0.001199408 Cytosol HSP90A 90A* Chaperonin 60* I2FKQ9 1.497301757 0.015466244 Mitochondrion CPN60C Chaperonin 11 A8J3C3 -1.921202539 0.018355322 Chloroplast; CPN11 mitochondria GrpE protein homolog Q945T2 -2.12638619 0.000523699 CGE1 Peptidylprolyl A8IW09 -2.177090798 0.004328963 FKB16-3 isomerase FKBP type* GrpE protein homolog Q945T1 -2.050517076 0.000844927 Mitochondrion CGE1 Amino acid metabolism S-adenosylmethionine A0A2K3D 2.16912801 0.019740384 Cytosol METM synthase LX7 Uncharacterized A0A2K3D 1.491765094 0.021405318 Cytosol CHLRE_0 protein KV5 7g344400v 5 Serine Q8W4V3 -1.587070898 0.001307894 SHMT hydroxymethyltransfer ase

149

Acetohydroxyacid A8IX80 -1.750425589 0.020400363 Chloroplast AAD1 dehydratase* LL-diaminopimelate A8IW39 -3.004098133 0.019129154 DPA1 aminotransferase Protein Biosynthesis Ribosomal protein L24 A0A2K3D 2.557933853 0.003746153 Cytosol RPL24 E61 40S ribosomal protein A8JGI9 2.329207377 0.019804299 Cytosol RPS7 S7 Elongation factor EF3 A8ISZ1 2.036689121 0.006317042 CHLRE_0 4g222700v 5 Ribosomal protein L3 A8ID84 1.883010232 0.008740299 Cytosol RPL3 Ribosomal protein A8I403 1.831163296 0.011590928 Cytosol RPS19 S19* 40S ribosomal protein A8J1G8 1.571491297 0.017705909 RPS6 S6* 40S ribosomal protein A8IMP6 1.425234188 0.012011262 RPS4 S4* Nop domain-containing A0A2K3D -1.383979032 0.00372515 Nucleus NOP56 protein 4E7 Plastid ribosomal A8IW44 -1.500751116 0.019051367 Chloroplast PRPL19 protein L19* Elongation factor G* A0A2K3E -1.550818484 0.008888525 Chloroplast EFG1 076 30S ribosomal protein Q08365 -1.631562351 0.018700606 Chloroplast RPS3 S3 Plastid ribosomal Q84U22 -1.654677446 0.005791956 Chloroplast PRPL4 protein L4*

150

Plastid ribosomal A8JE35 -1.668626833 0.005686262 Chloroplast PRPL3 protein L3* Uncharacterized A0A2K3D -1.723006151 0.001494541 Mitochondrion CHLRE_0 protein RT1 5g233800v 5 Uncharacterized A0A2K3C -2.28643668 0.011721623 Cytoplasm EFP1 protein* SJ6 Plastid ribosomal A8J5Y7 -2.520222566 0.009620662 Chloroplast RPS6 protein S6* H/ACA A0A2K3C -2.947586114 0.023499672 Nucleus CHLRE_1 ribonucleoprotein ZF8 3g567850v complex subunit 5 50S ribosomal protein Q8HTL3 -4.40991786 0.007147072 Chloroplast RPL23 L23 Cellular processes and transport NAD(P) A8JCP5 2.787355328 0.000694308 Mitochondrion CHLRE_0 transhydrogenase* 1g054500v 5 Uncharacterized A0A2K3C 1.943873727 0.00877991 PHC2 protein* XY7 Uncharacterized A0A2K3C 1.919340297 0.008973329 PHC13 protein* Y13 Histone 2A* A8HSB5 1.747852121 0.000113943 Nucleus HTA2 Histone 2A* Q42680 1.710207559 0.000522683 Nucleus HTA2 Histone 2A* A8IR81 1.710207559 0.000522683 Nucleus HTA20 Histone 2A* A8HWF6 1.710207559 0.000522683 Nucleus HTA10 Histone 2A* A8HWM8 1.710207559 0.000522683 Nucleus HTA11 HP domain-containing A0A2K3D 1.610936593 0.002883657 CHLRE_1 protein 4C8 2g524400v 5

151

Thioredoxin domain- A8JDA2 -1.827996818 0.011009963 CHLRE_1 containing protein* 7g715500v 5 38 kDa RNA-binding Q6EMK7 -2.311567171 0.004244904 Mitochondrion RB38 protein*

152

3.5 REFERENCES

Aguilera, A., Souza-Egipsy, V., & Amils, R. (2012). Photosynthesis in Extreme Environments. Artificial Photosynthesis. https://doi.org/10.5772/26940

Alexandratos, N., & Brunismas, J. (2012). WORLD AGRICULTURE TOWARDS 2030/2050. JESA Working Papers 12-03. https://doi.org/10.22004/ag.econ.288998

Allakhverdiev, S. I., & Murata, N. (2008). Salt stress inhibits photosystems II and I in cyanobacteria. Photosynthesis Research, 98(1–3), 529–539. https://doi.org/10.1007/s11120- 008-9334-x

Alric, J. (2010). Cyclic electron flow around photosystem I in unicellular green algae. Photosynthesis Research, 106(1–2), 47–56. https://doi.org/10.1007/s11120-010-9566-4

Alric, J. (2015). The plastoquinone pool, poised for cyclic electron flow? Frontiers in Plant Science, 6(JULY), 1–4. https://doi.org/10.3389/fpls.2015.00540

153

Cardona, T., Shao, S., & Nixon, P. J. (2018). Enhancing photosynthesis in plants: the light reactions. Essays in Biochemistry, 62(1), 85–94. https://doi.org/10.1042/EBC20170015

Dijkhuizen, L., & Harder, W. (1985). Current views on the regulation of autotrophic carbon dioxide fixation via the Calvin cycle in bacteria. Antonie van Leeuwenhoek, 50(1984), 473– 487.

Dolhi, J. M., Maxwell, D. P., & Morgan-Kiss, R. M. (2013). Review: The Antarctic

154

Chlamydomonas raudensis: An emerging model for cold adaptation of photosynthesis. Extremophiles, 17(5), 711–722. https://doi.org/10.1007/s00792-013-0571-3

Erickson, E., Wakao, S., & Niyogi, K. K. (2015). Light stress and photoprotection in Chlamydomonas reinhardtii. Plant Journal, 82(3), 449–465. https://doi.org/10.1111/tpj.12825

Finazzi, G., & Johnson, G. N. (2016). Cyclic electron flow: facts and hypotheses. Photosynthesis Research, 129(3), 227–230. https://doi.org/10.1007/s11120-016-0306-2

155

116(10), 4212–4217. https://doi.org/10.1073/pnas.1809812116

Govindjee, Shevela, D., & Björn, L. O. (2017). Evolution of the Z-scheme of photosynthesis: a perspective. Photosynthesis Research, 133(1–3), 5–15. https://doi.org/10.1007/s11120-016- 0333-z

Goyal, A., & Tolbert, N. E. (1990). Salicylhydroxamic acid (SHAM) inhibition of the dissolved inorganic carbon concentrating process in unicellular green algae. Plant Physiology, 92(3), 630–636. https://doi.org/10.1104/pp.92.3.630

He, Y., Fu, J., Yu, C., Wang, X., Jiang, Q., Hong, J., … Jiang, D. (2015). Increasing cyclic electron flow is related to Na+ sequestration into vacuoles for salt tolerance in soybean. Journal of

156

Experimental Botany, 66(21), 6877–6889. https://doi.org/10.1093/jxb/erv392

Huang, W., Zhang, S. B., Xu, J. C., & Liu, T. (2017). Plasticity in roles of cyclic electron flow around photosystem I at contrasting temperatures in the chilling-sensitive plant Calotropis gigantea. Environmental and Experimental Botany, 141(May), 145–153. https://doi.org/10.1016/j.envexpbot.2017.07.011

Huner, N. P. A., Öquist, G., & Sarhan, F. (1998). Energy balance and acclimation to light and cold,

157

1385(98).

Kato, Y., Ho, S. H., Vavricka, C. J., Chang, J. S., Hasunuma, T., & Kondo, A. (2017). Evolutionary engineering of salt-resistant Chlamydomonas sp. strains reveals salinity stress-activated starch-to-lipid biosynthesis switching. Bioresource Technology, 245, 1484–1490. https://doi.org/10.1016/j.biortech.2017.06.035

Kramer, D. M., & Evans, J. R. (2011). The importance of energy balance in improving

158

photosynthetic productivity. Plant Physiology, 155(1), 70–78. https://doi.org/10.1104/pp.110.166652

Laemmli, U. K. (1970). Cleavage of Structural Proteins during the Assembly of the Head of Bacteriophage T4. Nature, 227, 680–685.

Lemeille, S., & Rochaix, J. D. (2010). State transitions at the crossroad of thylakoid signalling pathways. Photosynthesis Research, 106(1–2), 33–46. https://doi.org/10.1007/s11120-010- 9538-8

Lima Neto, M. C., Cerqueira, J. V. A., da Cunha, J. R., Ribeiro, R. V., & Silveira, J. A. G. (2017).

159

Cyclic electron flow, NPQ and photorespiration are crucial for the establishment of young plants of Ricinus communis and Jatropha curcas exposed to drought. Plant Biology, 19(4), 650–659. https://doi.org/10.1111/plb.12573

Lucker, B., & Kramer, D. M. (2013). Regulation of cyclic electron flow in Chlamydomonas reinhardtii under fluctuating carbon availability. Photosynthesis Research, 117(1–3), 449– 459. https://doi.org/10.1007/s11120-013-9932-0

Melis, A. (1991). Dynamics of photosynthetic membrane composition and function. BBA - Bioenergetics, 1058(2), 87–106. https://doi.org/10.1016/S0005-2728(05)80225-7

Minagawa, J. (2016). A Supercomplex of Cytochrome bf and Photosystem I for Cyclic Electron Flow, 453–462. https://doi.org/10.1007/978-94-017-7481-9_23

Morgan-Kiss, R. M., Ivanov, A. G., Modla, S., Czymmek, K., Hüner, N. P. A., Priscu, J. C., …

160

Hanson, T. E. (2008). Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp., strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles, 12(5), 701–711. https://doi.org/10.1007/s00792-008-0176-4

Müller, P., Li, X. P., & Niyogi, K. K. (2001). Non-photochemical quenching. A response to excess light energy. Plant Physiology, 125(4), 1558–1566. https://doi.org/10.1104/pp.125.4.1558

161

Negrão, S., Schmöckel, S. M., & Tester, M. (2017). Evaluating physiological responses of plants to salinity stress. Annals of Botany, 119(1), 1–11. https://doi.org/10.1093/aob/mcw191

Oren, A. (2014). The ecology of Dunaliella in high-salt environments. Journal of Biological Research (Greece), 21(1), 1–8. https://doi.org/10.1186/s40709-014-0023-y

Priscu, J. C. (1998). Ecosystem Dynamics in a Polar Desert, 72.

Priscu, J. C., & Spigel, R. H. (1996). Evolution of temperature and salt structure of Lake Bonney, a chemically stratified Antarctic lake. Hydrobiologia, 321(3), 177–190.

Priscu, J. C., Wolf, F., Takacs, D., & Fritsen, H. (1999). Carbon Transformatiin a Perennially Ice- Covered Antarctic Lake, 49(12), 997–1008.

Raymond, J. A., & Morgan-Kiss, R. (2013). Separate Origins of Ice-Binding Proteins in Antarctic Chlamydomonas Species. PLoS ONE, 8(3), 6–11. https://doi.org/10.1371/journal.pone.0059186

162

Rochaix, J. D. (2007). Role of thylakoid protein kinases in photosynthetic acclimation. FEBS Letters, 581(15), 2768–2775. https://doi.org/10.1016/j.febslet.2007.04.038

Shetty, P., Gitau, M. M., & Maróti, G. (2019). Salinity Stress Responses and Adaptation Mechanisms in Eukaryotic Green Microalgae. Cells, 8(12), 1–16. https://doi.org/10.3390/cells8121657

Sithtisarn, S., Yokthongwattana, K., Mahong, B., Roytrakul, S., Paemanee, A., Phaonakrop, N., & Yokthongwattana, C. (2017). Comparative proteomic analysis of Chlamydomonas reinhardtii

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control and a salinity-tolerant strain revealed a differential protein expression pattern. Planta, 246(5), 843–856. https://doi.org/10.1007/s00425-017-2734-4

Sudhir, P., & Murthy, S. D. S. (2004). Effects of salt stress on basic processes of photosynthesis. Photosynthetica, 42(4), 481–486. https://doi.org/10.1007/S11099-005-0001-6

Szyszka-Mroz, B., Pittock, P., Ivanov, A. G., Lajoie, G., & Hüner, N. P. A. (2015). The antarctic psychrophile Chlamydomonas sp. UWO 241 preferentially phosphorylates a photosystem I-

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cytochrome b6/f supercomplex. Plant Physiology, 169(1), 717–736. https://doi.org/10.1104/pp.15.00625

Voss, I., Sunil, B., Scheibe, R., & Raghavendra, A. S. (2013). Emerging concept for the role of photorespiration as an important part of abiotic stress response. Plant Biology, 15(4), 713–

165

722. https://doi.org/10.1111/j.1438-8677.2012.00710.x

Ziotti, A. B. S., Silva, B. P., Sershen, & Lima Neto, M. C. (2019). Photorespiration is crucial for salinity acclimation in castor bean. Environmental and Experimental Botany, 167(May), 103845. https://doi.org/10.1016/j.envexpbot.2019.103845

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3.6 APPENDIX

Wt-LS Ev-LS 0.8 Wt-LS Ev-LS 0.8 Wt-HS Ev-HS B A Wt-HS Ev-HS

0.6 0.6 M

/F 0.4 0.4 V Y(PSII) F

0.2 0.2

0.0 0.0 Early-log Mid-log Late-log Early-log Mid-log Late-log

Wt-LS Ev-LS 1.0 Wt-HS Ev-HS 0.8 C D Wt-LS Ev-LS

Wt-HS Ev-HS 0.8 0.6

0.6 0.4 qL

0.4 Y(NPQ)

0.2 0.2

0.0 0.0 Early-log Mid-log Late-log Early-log Mid-log Late-log

Appendix Figure 3.1: PSII parameters for the Wt and Ev strain of C.reinhardtii under low and high salinity conditions during the growth curve. A. Maximum photosynthetic efficiency (Fv/Fm) B. Operating efficiency of PSII (YPSII). C. Proportion of open PSII (qL). D. Non-photochemical quenching (Y(NPQ)). Cells were far red adapted prior to the induction curve measurement to completely oxidize the photosynthetic pool. Actinic red-light intensity of 93 uE was used. Data are representative of three independent samples and the standard deviation are shown (n=3)

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20 A 5 B

4 15 g/ml) µ 3 10 2 5 Chlorophyll a/b 1 Chlorophyll (

0 0

Wt-LS Wt-HS Ev-LS Ev-HS Wt-LSWt-HSEv-LSEv-HS

Appendix Figure 3.2. Changes in chlorophyll content and ratio of wildtype and evolved strain of C. reinhardtii under low (LS) and high salinity (HS). A. Total chlorophyll content (n=3, ±SD). B. Chlorophyll a/b ratio shown as a violin plot, means are depicted as broken lines (n=3).

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Chapter 4

Impact of salinity-tolerance versus-acclimation on the structure and function of the photochemical apparatus: a comparative study

Isha Kalra, Xin Wang, Rachael Morgan-Kiss

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Chapter IV. Impact of salinity-tolerance versus-acclimation on the structure and

function of the photochemical apparatus: a comparative study

4.1 INTRODUCTION

Photosynthesis consists of linear electron flow (LEF) that transfers electrons from photosystem II

(PSII) to photosystem I (PSI) via cytochrome b6f and mobile electron carriers. LEF produces 2.57 ATP and 2 NADPH per electron. However, the downstream carbon fixation requires 3 ATP

and 2 NADPH for assimilation of 1 CO2 molecule in carbon fixation (Kramer & Evans, 2011).

Cyclic electron flow (CEF) involves electron transfer from PSI to plastoquinone (PQ) to cyt b6f and back to PSI via plastocyanin (PC). As the electrons are cycling back to PSI, CEF only produces ATP via acidification of lumen by cyt b6f and no NADPH is produced (Yamori & Shikanai, 2016). Thus, the ATP produced from CEF mitigates the imbalance between photosynthesis and carbon fixation. When light absorbed is greater than needed for growth and carbon fixation it leads to energy imbalance and over-reduction of the plastoquinone pool, which can damage the photosynthetic apparatus by formation of reactive oxygen species (ROS). Photosynthetic organisms maintain ‘photostasis’, where the amount of energy absorbed is relatively balanced with the amount needed to drive downstream metabolism (Ensminger et al., 2006). Many environmental stresses can cause imbalances in the energy demands and increase the risk of ROS production. In their natural environment, organisms must respond to various levels of environmental stress (including high light, low temperatures, high salinity, nutrient deficiency), which may last for a few minutes (short-term or transient) or persist for days to years (long-term) (Kono & Terashima, 2014). Most studies that focus on understanding the role of CEF during stress, focus on its involvement under transient stress conditions. Indeed, CEF induction under transient stress has often involved extreme treatments, such as dark/anoxic treatment or incubation in nitrogen-free growth medium. Thus, CEF is generally considered to be involved in transient stress acclimation; however, its role in long-term stress acclimation has not been thoroughly studies. While the CEF mechanism is not fully understood, formation of protein ‘supercomplexes’ appear to play an important role in the activation of CEF (Minagawa, 2016). In

170 the last decade, the contribution of PSI-supercomplexes in initiating CEF is has been extensively studied in model organisms such as C. reinhardtii (Minagawa, 2016). As the role of CEF is mostly studied in the context of transient stress, similarly, the presence of supercomplexes has also been studied under these short-term conditions in C. reinhardtii. In particular, researchers have isolated the supercomplex after either exposure to inhibitors that lock the organism is state 2 or under anaerobic conditions that also leads to state 2 transition (Iwai et al., 2010; Takahashi et al., 2016; Terashima et al., 2012). Thus, the supercomplexes are often studied and characterized in transient and sometimes even physiologically irrelevant conditions. Salinity stress is one of the major abiotic stresses that plants and other photosynthetic organisms are exposed to in their natural habitats, reducing productivity (Morton et al., 2019). The excess sodium ions can build up in the cytoplasm causing ion toxicity and also disturb the osmotic balance (Kumar et al., 2018). Photosynthetic organisms respond to salinity stress in two major ways: (i) accumulation of compatible osmolytes such as glycerol in the cell to maintain osmotic balance, and (ii) active exclusion of sodium ions to maintain ion homeostasis inside the cell (Goyal, 2007a; He et al., 2015). These two processes are excess ATP consuming processes, requiring organisms that are dealing with salinity to continuously keep up with the energy demands. Photoautotrophic organisms must use photosynthesis mediated ATP production, and cannot rely on external sources of energy. CEF is one of the major pathways by which photosynthetic organisms fine tune the ATP/NADPH ratio based on downstream energetic demands (Suorsa, 2015). Although, significant research has been focused on short-term salinity stress response in plants and algae (Neelam & Subramanyam, 2013; Sudhir & Murthy, 2004; N. Wang et al., 2018), few studies have looked into long-term salinity acclimatory response in green algae (Shetty et al., 2019). These few studies have focused on characterizing the metabolic changes associated with salinity acclimation using transcriptomics, proteomics and other metabolic measurements in C. reinhardtii (Perrineau et al., 2014; Sithtisarn et al., 2017). However, potential upstream changes associated with the reorganization of photosynthetic electron transport have been largely ignored. The paucity of research considering salt-induced changes to the photosynthetic apparatus is a symptom of the typical design of salinity acclimation experiments. Most past experiments were performed in the presence of alternate carbon source, acetate. Under these conditions, the photoheterotrophic C. reinhardtii bypasses photosynthesis and predominantly uses acetate as a

171 source of energy (Perrineau et al., 2014). As a result, it is important to conduct salinity acclimation experiments in strict phototrophic conditions where the role of photosynthesis electron transport in salinity acclimation can be fully dissected. Growth under excessively high salinity conditions is one of the stressful conditions: few model photosynthetic organisms exhibit elevated salinity tolerance. One exception is the model halophile, Dunaliella, a green alga which if often the dominant primary producer in hypersaline environments (Oren, 2014). Comparative studies between the genetic model C. reinhardtii and Dunaliella spp. is difficult, as the genera are distantly related. One of the few naturally salinity tolerant Chlamydomonas studied extensively is the Antarctic Chlamydomonas sp. UWO241. UWO241 is a strict photoautotroph that has been shown to utilize increased CEF during acclimation to long-term high salinity. Recent work has shown that elevated CEF in this halotolerant psychrophile contributes to both PSI and PSII photoprotection and production of extra ATP, the latter is utilized to balance downstream metabolic needs (Kalra et al., 2020). While it is often assumed that global climate change is mainly associated with increasing temperatures, this is a simplistic view. In fact, the direct and indirect effects of environmental change on the growth and productivity of photosynthetic organisms residing in different habitats is complex. As the climate change exacerbates, there is a growing need for an improved understanding of how organisms will respond to and survive a myriad of stress conditions, especially long-term steady-state stresses (Alexandratos & Brunismas, 2012). The function of the photosynthetic apparatus is key to survival under environmental change: the role of CEF is an essential pathway in all photosynthetic organisms, making it an ideal candidate to study stress acclimation in the context of climate change (Kramer & Evans, 2011). Understanding how CEF can help plant and algal survival under physiologically relevant, steady-state stress conditions can help us engineer photosynthetic organisms to better withstand climate change in the future. Most photosynthetic research has intensively focused on a handful of model organisms under controlled laboratory conditions. These approaches offer a variety of advantages; however, there is a significant knowledge gap in identifying natural mechanisms which help photosynthetic organisms deal with stress in their environment. The lack of studies in native photoautotrophs is important for two major reasons: i) first, there may be novel physiological strategies which have been overlooked, and ii) second, organisms from different habitats could exhibit different sensitivities to environmental change. Non-model organisms that are adapted to very long-term

172 stress lasting 100s-1000s of years represent under-exploited reservoirs of novel adaptive mechanisms (Dolhi et al., 2013). The Antarctic psychrophile Chlamydomonas sp. UWO241 was isolated from deep photic zone of the ice-covered lake Bonney in the McMurdo Dry Valleys (Morgan et al., 1998). Extensive studies on this organism have demonstrated that it is a psychrophilic, halotolerant green alga that has reorganized its photosynthetic apparatus to downregulate short-term acclamatory strategies like state transitions but maintain strong steady-state levels of CEF (R. Morgan-Kiss et al., 2002; Morgan et al., 1998; Szyszka et al., 2007). A recent study by Szyszka et. al demonstrated that under high salinity stress, UWO241 forms a thylakoid membrane protein supercomplex composed of PSI, cyt b6f, PGR5 (Proton gradient 5), LHCI (Light Harvesting Complex I), as well as two novel proteins, FtsH and PsbP. Phosphorylation of the latter proteins (FtsH and PsbP) appears to be essential for supercomplex stabilization. Formation of the UWO241 complex is distinct from that described in other model organisms (ie. C. reinhardtii) as it is independent of state transitions. Since C. sp. UWO241 exhibits several novel strategies to survive long-term stress, the question of whether other extremophile exhibit comparable strategies was posed. A second photopsychrophile, Chlamydomonas sp. ICE-MDV was recently isolated from Lake Bonney and is the dominant alga in the upper, freshwater layers of the lake (W. Li & Morgan-Kiss, 2019; W. Li, Podar, & Morgan-Kiss, 2016). ICE-MDV is also cold-adapted, requiring low temperatures for growth; however, unlike UWO241, it dominates relatively freshwater depths of Lake Bonney. Thus, these sister Chlamydomonas species provide a unique comparative opportunity to further explore photosynthetic adaptation to low temperature and high salinity. Our previous work showed that acclimation to high salinity is required for assembly of the supercomplex formation and maximal CEF rates in UWO241. Here, we hypothesized that CEF and supercomplexes are important in long-term salinity stress acclimation in other Chlamydomonas species. As a consequence of adaptation to permanent low temperature/high salinity stress, UWO241 has lost major short-term responses, most notably state transitions. We hypothesized that acclimation to long-term salinity stress will inhibit or attenuate the capacity for state transitions. To test these hypotheses, we conducted a comparative study of three different Chlamydomonas species exhibiting a gradient of natural salinity tolerance: two Antarctic algae with different salt sensitivities, C. sp. UWO241 and C. sp. ICE-MDV, and the salt-sensitive

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model mesophile C. reinhardtii. We determined the maximum salinity tolerance of each Chlamydomonas strain, and then tested the ability of each organism to assemble a supercomplex and modulate CEF rates. Last, state transition capacity across all strains was compared under low and high salinity conditions.

4.2 METHODS:

4.2.1 Culture conditions, growth physiology.

Three different Chlamydomonas species were used in this study: C. sp. UWO241 (UWO241; CCMP1619), C. sp. ICE-MDV (ICE-MDV) and the model C. reinhardtii (UTEX 90). All three species were first grown in Bold’s Basal Media (BBM, 0.43 mM NaCl) (Low salt, LS). Based on previous studies, UWO241 and ICE-MDV cultures were grown under a temperature/irradiance regime of 8oC/50 photons μmol m-2s-1 (Cook et al., 2019; Morgan et al., 1998). C.reinhardtii UTEX 90 was grown in BBM (Low Salt, LS) at 20°C/100 μmol photons m- 2s-1. All cultures were grown in 250 ml glass pyrex tubes in temperature regulated aquaria under a 24 h light cycle and were continuously aerated with sterile air supplied by aquarium pumps (Morgan-Kiss, Ivanov, & Huner, 2002). For salinity tolerance study, cultures were grown in increasing concentration of NaCl supplemented BBM (0-700 mM NaCl for UWO241 and ICE-MDV, 0-200 mM NaCl for C. reinhardtii). Growth was monitored daily by optical density at wavelength of 750 nm. Maximum growth rates were calculated using natural log transformation of the optical density values during the exponential phase. Three biological replicates were performed. For salinity stress acclimation, cultures were grown in maximum tolerated salinity levels and sub-cultured after reaching log-phase, at least 2-3 times. All subsequent experiments were conducted on high salinity acclimated (High salt, HS) log-phase cultures.

4.2.2 State transition induction

State transition experiments were conducted on both low and high salinity acclimation cultures. Cultures were harvested in the mid-log phase and induced in either state 1 or state 2 through addition of chemical inhibitors as described before (Iwai et al., 2010). Briefly, for state 1 induction, mid-log phase cells were incubated in 10 µM DCMU to completely oxidize the PQ

174

pool. For state 2 induction, cells were incubated in 5 µM FCCP for 20 min. State transition response was measured through either 77K spectra or PSII fluorescence as described below.

4.2.3 Low temperature (77K) fluorescence spectra.

Low temperature Chl a fluorescence emission spectra of whole cells and isolated Chl-protein complexes were measured using a Luminescence Spectrometer LS50B (Perkin Elmer, USA) as described in Morgan et al. (2008) at an excitation wavelength of 436 nm, using slit widths of 5 (isolate complexes) or 8 nm (whole cells) (R. M. Morgan-Kiss et al., 2008). Prior to the measurement, cultures were dark adapted for 10 min. Decompositional analysis was performed using a non-linear least squares algorithm using Microcal OriginPro Version 8.5.1 (Microcal Origin Northampton, MA). The fitting parameters for the Gaussian components (position, area and full width half-maximum, FWHM) were free running parameters.

4.2.4 PSII fluorescence state transition measurement

Room temperature PSII fluorescence measurements were done on cultures induced in state 1 or state 2 as described above. 2 ml of cultures from log-phase were used for measurements.

Measuring light was switched on in the dark and minimal PSII fluorescence (FO) was measured. Then cultures were exposed to 200 μmol photons m-2s-1 of actinic red light (λmax=620 nm, 10 −2 Wm , Scott filter RG 715) to measure maximum fluorescence (FM). State transition capacity ST1 ST2 ST1 was calculated using FM values under state 1 and state 2 and the formula: (FM - FM )/ FM

ST1 ST2 %, to show percent state transition capacity (Girolomoni et al., 2017), where FM and FM are the maximal PSII fluorescence under state 1 and 2 respectively.

4.2.5 SDS-PAGE and Immunoblotting.

SDS-PAGE was performed using Bio-Rad Mini-Protean system and 12% Urea-SDS gel (Laemmli, 1970). Thylakoid membranes were denatured using 50 mM DTT and incubated at 70°C for 5 min. Samples were loaded on equal protein basis (10 μg total protein). Proteins were transferred to nitrocellulose membranes using cold-wet transfer at 100 V for 2.5 hours. The membrane was blocked with TBST (Tris Buffer Saline Tween) buffer with 5% milk (Carnation). A primary antibody against P-Thr (Catalog # MA5-27976, Thermo Fisher) was used at 1:500 dilution to probe for phosphorylated threonine sites. Membranes were then exposed to Protein A

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conjugated to horseradish peroxidase and blots were detected with ECL SelectTM Western Blotting Detection Reagent (Amersham).

4.2.6 P700 oxidation-reduction kinetics

Actinic red light induced photooxidation of P700 was used to determine rates of CEF as previously described (Alric, Lavergne, & Rappaport, 2010; R. M. Morgan-Kiss et al., 2002). A volume of exponential phase cultures representing 25 μg Chl a was dark adapted for 10 min in presence of DCMU to block LEF and then filtered onto 25 mm GF/C filters (Whatman). Filters were measured on the Dual-PAM 100 instrument using the leaf attachment. The proportion of photooxidizable P700 was determined by monitoring absorbance changes at 820 nm and expressed as the parameter (∆A820/A820). The signal was balanced, and the measuring light switched on. Actinic red light (λmax=620 nm, 10 Wm−2, Scott filter RG 715) was then switched on to oxidize P700. After steady-state oxidation levels were reached, the AL was switched off to

+ red re-reduce P700. The half time for the reduction of P700 to P700 (t½ ) was calculated after the AL was turned off as an estimate of relative rates of PSI-driven CEF (Ivanov et al., 1998). The re-reduction time for P700 was calculated using MicrocalTM OriginTM software (Microcal Software Inc., Northampton, MA, USA).

4.2.7 Supercomplex isolation.

Sucrose step density centrifugation was used to isolate supercomplexes from exponentially grown cultures according to Szyszka-Mroz et al. (2015) with some modifications (Szyszka-Mroz et al., 2015). Every step was performed in darkness and on ice. All buffers contained phosphatase (20 mM NaF) and protease (1 mM Pefabloc SC) inhibitor. Cells were collected by centrifugation and the pellet was washed twice in Buffer 1 (0.3 M Sucrose, 25 mM Hepes-KOH

[pH 7.5], 1mM MgCl2). Cells were disrupted using French press, as described above and broken cells were spun down at 50,000g for 30 min. The pellet was resuspended in Buffer 2 (0.3 M Sucrose, 5 mM Hepes-KOH [pH 7.5], 10 mM EDTA) and centrifuged at 50,000 x g for 30 min. The thylakoid pellet was resuspended gently in Buffer 3 (1.8 M Sucrose, 5mM Hepes-KOH [pH 7.5], 10 mM EDTA) and transferred to Ultra-clear tube (Catalogue No., 344060, Beckman Coulter, USA). The thylakoid fraction was overlayed with Buffer 4 (1.3 M Sucrose, 5mM Hepes-KOH [pH 7.5], 10 mM EDTA) followed by Buffer 5 (0.5 M Sucrose, 5mM Hepes-KOH [pH 7.5]). This sucrose step gradient was ultra-centrifuged at 288,000g for 1 hour at 4°C using

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Sw40Ti rotor (Beckman coulter, USA). Purified thylakoids were collected and diluted (3-fold) in Buffer 6 (5 mM Hepes-KOH [pH 7.5], 10 mM EDTA) and centrifuged at 50,000xg for 20 minutes to pellet the membrane. Linear sucrose gradients were made using freeze thaw method with Buffer 7a (1.3 M Sucrose, 5 mM Hepes-KOH [pH 7.5], 0.05% ß-DDM) and Buffer 7b (0.1 M Sucrose, 5 mM Hepes-KOH [pH 7.5], 0.05% ß-DDM). Briefly, two dilutions of Buffers 7a and 7b were made, Buffer 7-1 (2x Buffer 7a + 1x Buffer 7b) and Buffer 7-2 (1x Buffer 7 a + 2x Buffer 7b). To make the gradient, first 3 ml of Buffer 7a was poured into 12 ml ultra-clear tubes followed by flash freezing in liquid nitrogen. Next, Buffer 7-1 was poured on top, followed by flash freezing. This was repeated for Buffer 7-2 and Buffer 7b respectively. The frozen gradients were kept at 4°C overnight to thaw. For supercomplex isolation, thylakoid membranes (0.4 mg Chl) were resuspended in 1% n-dodecyl-beta-maltoside (ß-DDM) (Catalogue number D99020, Glycon Biochemicals, Germany) and incubated on ice in the dark for 25 min. Membranes were spun down to remove insoluble material and loaded onto a linear sucrose gradient described above (0.1 – 1.3 M sucrose) containing 0.05% ß-DDM. Gradients were centrifuged at 288,000g for 21 hours at 4°C using SW40Ti rotor (Beckman Coulter, USA). Protein complexes were extracted using a 21-gauge needle.

4.2.8 Sample preparation for proteomics.

For identifying protein components in the supercomplex, the complex was harvested and 30 μg of total protein was processed for proteomics following the previously published method by Wang et al. (Wang et al., 2016). Specifically, samples were treated with 8 M Urea/5 mM DTT for 1 hour at 37°C, followed by alkylation with 15 mM iodoacetamide in dark for 30 minutes at room temperature. Samples were then diluted 4- folds with 50 mM Tris-HCl buffer and digested using Mass-spectrometry Grade Trypsin Gold (Promega, Madison, WI) at 1:100 w/w concentration for 16.5 hours at 37°C. The digested samples were cleaned using Sep-Pak C18 plus desalting columns (Waters Corporation, Milford, MA).

4.2.9 Proteomic analyses by liquid chromatography-tandem mass spectrometry (LC-MS/MS)

The digested peptides were directly loaded onto a capillary C18 column without fractionation, and further analyzed in a two dimensional (2D)-LC-MS/MS Thermo LTQ Orbitrap XL mass spectrometer. The full mass spectra were recorded in the range of 350-1800 m/z with the resolution of 30,000. The top 12 peaks of each scan were selected for MS/MS analysis. The

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MS/MS raw data was analyzed by first converting into MS2 files, followed by database search using ProLuCID (Xu et al., 2006). The UWO241 protein database was generated based on our transcriptomics data supplemented with 37 common contaminants, and their reversed sequences as quality control system to restrain false positive discovery to 0.05. C. reinhardtii protein database was downloaded from NCBI containing both Swiss-Prot and TrEMBL entries. Proteins were identified and analyzed using PatternLab for Proteomics (Carvalho et al., 2016).

4.3 RESULTS

4.3.1 Salinity tolerance the three Chlamydomonas species

To identify the salinity sensitivity for all three Chlamydomonas species, we conducted a salinity gradient growth experiment. Based on the native habitat of the Antarctic species (ie. the hypersaline Lake Bonney) as well as previous literature on salt tolerance in UWO241 (Pocock et al. 2010) and C. reinhardtii (Subramanyam et al. 2010), we selected the following salinity concentrations: (i) ICE-MDV and UWO241 were grown in salinity concentrations of 0 to 700 mM NaCl (salinity concentration at 17 m depth of Lake Bonney), and (ii) C. reinhardtii was grown in 0.43 - 200 mM NaCl (Subramanyam et al. 2010). (Fig. 4.1). As expected, the halotolerant UWO241 grew well under the full range of salinity conditions. Despite its native environment of hypersalinity, UWO241 had the shortest lag phase and a doubling time of 72 ± 2.44 hrs (Fig. 4.1 A). Growth under the moderate salinity stress of 250 or 500 mM caused a ~1.6-fold increase (p < 0.05) in doubling time (113 ± 24.13 and 119 ± 3 hrs, respectively). Last, while the lag phase was significantly longer in UWO241 cultures grown in 700 mM NaCl, once acclimated, these cultures exhibited the fastest doubling time (55 ± 3.38 hr), 1.3-fold faster relative to BBM-grown cultures. ICE-MDV was more recently isolated from Lake Bonney, where it dominates the shallow, freshwater depths (Li et al. 2016; Li and Morgan-Kiss 2019). In the salinity gradient experiment, ICE-MDV grew fastest under control conditions (0.43 mM NaCl, doubling time of 82 ± 0.7 hr), but exhibited the ability to grow well under a salinity regime of either 250 mM or 500 mM NaCl (doubling times of 98.9 ± 10.6 and 115.2 ± 11.2 hrs, respectively) (Fig. 4.1 B). However, unlike UWO241, which has a robust growth at 700 mM NaCl, ICE-MDV was unable to grow in 700 mM NaCl.

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The model mesophile C. reinhardtii had the highest growth rate under control conditions (doubling time of 55.6 hrs), followed by 50 mM NaCl (doubling time of 68.2 hrs) (Fig. 4.1 C). For C. reinhardtii, 100 mM NaCl was the maximum salinity that the cultures exhibited some growth; however, the cultures failed to grow beyond an OD750 of 0.6. Last, after a few days of slight growth in the upper salinity levels of 150 mM and 200 mM NaCl, C. reinhardtii failed to grow further and entered death phase in both of these salinity treatments. Based on these growth physiology results, we chose the following salinity levels for further experiments testing long- and short-term acclimation responses. For low salinity (LS), all strains were grown in BBM medium (0.43 mM NaCl). For high salinity (HS), we used BBM supplemented with, (i) 700 mM NaCl for UWO241, (ii) 500 mM NaCl for ICE-MDV and (iii) 50 mM NaCl for C. reinhardtii. Cultures were acclimated by serial sub-culturing in the same condition for 14 – 30 days depending upon the growth rate.

4.3.2 Photosystem I activity

Our previous work on the halotolerant UWO241 showed that this organism exhibits higher CEF rates under a growth environment of high salinity (Kalra et al., 2020). However, not many studies have focused on the role of CEF in long-term stress acclimation. To further expand our knowledge on the ubiquity of sustained CEF under long-term environmental stress, we monitored PSI activity in the sister psychrophilic Antarctic algae ICE-MDV and the model green alga C. reinhardtii, which showed moderate and minimal salinity tolerance (Fig. 4.1). P700 oxidation/reduction kinetics were conducted to monitor changes in P700 photooxidation activity and PSI-mediated CEF in response to long-term salinity acclimation. Prior to the measurement, cultures were incubated with DCMU to block electron transport from PSII to PSI. Cultures are exposed to red actinic light (AL), leading to oxidation of P700 to P700+, and the absorbance changes was be monitored at 820 nm. After 10 s of illumination, a

red steady state of A820 was reached and the AL was switched off. Dark re-reduction rate (t½ ) of

+ P700 was measured post AL illumination (Appendix Fig. 4.1), Under these conditions (ie. in the presence of the PSII inhibitor, DCMU), reduction of P700 is due to alternative electron transport pathways, mainly PSI mediated CEF (Ravanel et al, 1994, Ivanov et al, 2012). LS-grown

red cultures of UWO241 exhibited a t½ of 258.8 ± 44.7 ms. In agreement with previous literature (Kalra et al. 2020; Szyska-Mroz et al. 2016) acclimation of UWO241 to HS resulted in a 1.6-fold

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red faster re-reduction rate (mean t½ = 162.2 ± 13.6 ms) (Fig. 4.2 A). Under LS, ICE-MDV

red red exhibited a comparable t½ value as LS-grown UWO241 (mean t½ = 291.2 ± 41.94 ms). ICE-

red red MDV responded to HS by a 2.3-fold faster t½ (mean t½ = 124 ± 31.5 ms). On the other hand, C.

red red reinhardtii exhibited a slower t½ rate in LS media (mean t½ = 494.9 ± 42.724 ms) which was around 1.6 - 1.9-fold slower than LS-grown ICE-MDV and UWO241. However, similar to the psychrophiles, C. reinhardtii displayed 1.5 times faster re-reduction rates after acclimation to

red salinity stress (mean t½ = 311.1 ± 10.362 ms) (Fig. 4.4 C).

We also measured change in P700 absorbance after AL illumination which reflects the + redox state of P700. The change in absorbance of P700 radical after excitation of PSI by AL was

measured as ∆A820 /A820. Both the psychrophiles, UWO241 and ICE-MDV (Fig 4.2 D, E),

displayed lower ∆A820 /A820 values when compared to C. reinhardtii (Fig 4.2 F) (1.5 and 1.7 times lower, respectively), indicating a reduced capacity for PSI oxidation. Interestingly, the

∆A820 /A820 values were further reduced significantly in the psychrophiles (Fig 4.2 D, E) after salinity stress acclimation, while C. reinhardtii did not show any significant change in ∆A820

/A820 after salinity stress acclimation (Fig. 4.2 F). Thus, all three strains responded to HS by increasing CEF; although, both psychrophiles exhibited constitutively higher CEF relative to the mesophile.

4.3.3 Effect of long-term stress high salinity acclimation on short-term state transition response

State transitions are a universal which has been documented in taxonomically diverse taxa as a short-term response to overexcitation of the photosynthetic electron transport chain (Rochaix, 2007). A state 1 to 2 transition is manifested as a transfer in absorbed light energy from PSII to PSI, mediated by migration of a sub-population of LHCII. Previous research showed that UWO241 is a natural state transition mutant (Morgan-Kiss et al. 2002); however, the capacity for state transitions in the sister species ICE-MDV has not been tested. To this end, we conducted state transition tests on all three Chlamydomonas species after low and high salinity acclimation. Chemical inhibitors DCMU and FCCP were used to artificially induce state 1 and state 2 conditions respectively. The state transition ability was tested by using low temperature 77K fluorescence spectra as well as room temperature maximum PSII fluorescence.

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PSI fluorescence is minimal under room-temperature and 77K fluorescence spectra analysis overcomes this problem by keeping the sample at low temperature of 77 Kelvin. At this temperature, both PSII and PSI Chl a fluorescence can be measured using a excitation wavelength of 436 nm. This produces fluorescence emission spectra, where PSII fluorescence peak is at 682-685 nm whereas PSI peak is at 712-715 nm. During state 1, the plastoquinone pool is completely oxidized and LHCII remains attached to PSII. This phenomenon can be observed as higher PSII fluorescence relative to PSI (Fig. 4.3). On the other hand, PSII fluorescence is reduced under conditions triggering a transition from state 1 to state 2. As expected, under both low and salinity condition, PSI fluorescence emission yields were very low. Although PSI peak is low under both salinity levels, UWO241 exhibited a markedly lower PSI fluorescence yield under high salinity. Furthermore, induction of a state transition had no significant effect on PSI fluorescence in UWO241 grown under either condition (Fig. 4.3 A). While ICE-MDV was isolated from the same Antarctic lake as UWO241, its 77K fluorescence emission spectra characteristics were markedly different from that of UWO241 (Fig. 4.3B). First, under low salinity, ICE-MDV exhibited detectable levels of PSI (Fig. 4.3 B). In addition, when exposed to state 2 conditions, LS-grown ICE-MDV responded with a 1.25-fold increase in PSI fluorescence, suggesting that unlike UWO241, ICE-MDV has retained the ability to undergo state transitions (Fig. 4.3 B). Acclimation to HS resulted in a loss of PSI fluorescence in ICE-MDV, resulting in a 1.2-fold reduction in HS- versus LS-conditions. Last, high salinity acclimation in ICE-MDV had a pronounced effect on its capacity for state transitions. Under state transition conditions, no detectable change in PSI fluorescence was observed in HS- acclimated ICE-MDV cells (Fig. 4.3 B). In agreement with previous literature, under low salinity C.reinhardtii exhibited a typical fluorescence spectrum and the ability to undergo a state transition (Fig. 4.3 C, black line)., Growth under high salinity resulted in a 1.2-fold decrease in PSI fluorescence yield (Fig. 4.3 C, red line). Last, unlike the psychrophiles, HS-acclimated C. reinhardtii cultures retained the ability to undergo a state transition, exhibiting a comparable response to state 1 conditions as the LS-grown cells (Fig. 4.3 red line). State transition capacity can also be measured as a change in in maximum fluorescence of

PSII (FM) at room temperature (Appendix Fig. 4.2). We used this measurement to validate the ST1 ST2 ST1 77K fluorescence emission data. We used the formula: (FM - FM )/ FM %, to show percent

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ST1 ST2 state transition capacity (Girolomoni et al., 2017), where FM and FM are the maximal PSII fluorescence under state 1 and 2, respectively. If a state transition occurs, one would expect to detect a positive value; however, for cultures of either LS- or HS-UWO241, the calculated state transition capacity was negative, confirming the 77K fluorescence results indicating that UWO241 has little to no capacity for state transitions under either growth conditions (Fig. 4.3 D). Similar negative values have been previously seen for stt7 mutants in C.reinhardtii (Girolomoni et al., 2017). In contrast, both LS-grown ICE-MDV and C. reinhardtii cells exhibited 36 and 29 % state transition capacity, which also agreed with our 77K fluorescence emission findings. Interestingly, both ICE-MDV and C. reinhardtii exhibited a significant reduction in state transition capacity, following acclimation to high salt (3.5- and 2.2-fold, respectively, relative to LS conditions; p<0.01; Fig. 4.3 E, F). These results differed from that of the 77K fluorescence emission spectra, which showed minimal effect of high salinity on the state transition capacity of either algal species.

4.3.4 Thylakoid protein phosphorylation

In C. reinhardtii, several key photosynthetic proteins are phosphorylated in response to changes in environmental conditions. State transitions are accompanied by transient phosphorylation of LHCII proteins through stt7 kinase (Lemeille & Rochaix, 2010). Previous reports have shown that the thylakoid proteome of UWO241 is under-phosphorylated relative to other photosynthetic organisms and exhibited novel high molecular weight phospho-proteins (Morgan-Kiss et al. 2002; Szyszka et al. 2007). We compare the phospho-protein profiles of the three strains acclimated to either LS or HS (Fig. 4.4). When probed with a phospho-threonine antibody, phosphorylate of major LHCII proteins was not observed in UWO241 grown under either LS or HS conditions. (Fig 4.4, left panel). On the contrary, UWO241 exhibited phosphorylation of several high molecular weight proteins (~150 and 250 kDa) under both LS and HS. Our state transition tests showed that unlike UWO241, ICE-MDV exhibits a typical state transition capacity. Since state transitions are dependent upon reversible phosphorylation of major LHCII, we probed thylakoids of ICE-MDV grown under LS and HS to see whether LHCII proteins are also phosphorylated. In agreement with the state transition results, ICE-MDV exhibited phosphorylation of some LHCII proteins (type III and IV) under both LS and HS condition; however, some phospho-LHCII proteins that were detected in C. reinhardtii were not

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detectable (Fig. 4.4, middle panel). Remarkably, thylakoids isolated from LS- or HS-grown ICE-MDV also exhibited phosphorylation of several high molecular weight proteins (~150 kDa), albeit at lower level compared with UWO241. Thus, the two psychrophiles share this interesting phenomenon of large, unknown phosphoproteins in the thylakoid membranes. As expected, C. reinhardtii exhibited phosphorylation of several thylakoid proteins, specifically phosphorylated LHCII (type I, II, III and IV) (Fig. 4.3, right panel). Growth in high salinity did not alter the pattern or abundance of phosphoproteins of C. reinhardtii. Last, in contrast with the Lake Bonney algae, we did not detect phosphorylation of larger proteins in C. reinhardtii under either growth condition.

4.3.5 Assembly of protein supercomplexes under high salinity

PSI-supercomplexes have been shown to be associated with conditions promoting CEF. In C. reinhardtii, supercomplexes have been isolated from stress conditions such as state 2 transition and anoxia. These supercomplexes are shown to contain PSI, LHCI and LHCII proteins, as well

as other proteins such as CAS (Calcium associated protein) and PETO (cytb6f subunit) under certain conditions. UWO241 is one of the first Chlamydomonas species shown to possess a supercomplex under a long-term stress, ie. high salinity (Szyszka et al., 2015). Formation of the UWO241 supercomplex is proposed to be essential for maintaining sustained high rates of CEF (Kalra et al. 2020) As C. reinhardtii and ICE-MDV also increase CEF during salinity stress acclimation (Fig. 4.2), we hypothesized HS acclimation may also be associated with formation of PSI supercomplex in C. reinhardtii and ICE-MDV. To identify and isolate the PSI- supercomplex, we conducted sucrose density gradient centrifugation on solubilized thylakoid membranes from the three Chlamydomonas species after acclimation to salinity stress. As a comparison, we also ran sucrose density gradients on thylakoids from C. reinhardtii exposed to either state 1 and state 2, classic short-term conditions for inducing supercomplex formation in this organism. Sucrose density gradient centrifugation on solubilized thylakoid membranes resolves several major chlorophyll-binding protein complexes. Under nonstress conditions, there are typically three major pigment-containing bands: (i) trimeric LHCII (band 1), (ii) PSII core (band 2) and (iii) PSI-LHCI complex (band 3). Separation of these three complexes is shown in Figure 4.5A. Under State 1, thylakoid of LS-grown C. reinhardtii exhibited all three bands. In contrast,

183 when LS-grown cells are exposed to State 2 conditions, a fourth, heavier band appears, below the PSI-LHCI band, representing the PSI supercomplex (Fig. 4.5 A, band 4). Next, we compared the banding patterns in sucrose density gradients from thylakoids isolated from all three Chlamydomonas species after acclimation to salinity. In response to HS, all strains exhibited a reduction in the relative levels of PSI (Band 3). In addition, all HS- acclimated samples from all three Chlamydomonas species exhibited the appearance of the supercomplex band (band 4; Fig. 4.5A, B). However, there were some distinct differences in the banding patterns. For example, the LHCII trimer migrated lower in both ICE-MDV and UWO241 compared to C. reinhardtii, suggesting that the major LHCII antenna are larger in both psychrophiles. Bands from all gradients were collected for 77K Chl fluorescence analysis (Fig. 4.5 C-G). In C. reinhardtii exposed to short-term conditions, emission peaks for the major Chl-protein complexes agreed with previous reports. In state 1, Band 1 showed a fluorescence peak at 678 nm, coinciding with LHCII whereas Band 2 fluoresced at 682 nm indicating PSII. Band 3 exhibited emission peaks at both 678 and 712, indicating presence of PSI and LHCs (Fig. 4.5 C). In state 2, Band 1-3 coincided with LHCII, PSII and PSI fluorescence peaks respectively, and Band 4 had peaks at both PSI and LHCII fluorescence maxima (Fig. 4.5 D). Bands 1 and 2 from HS-acclimated C. reinhardtii exhibited typical emission peaks associated with LHCII and PSII, respectively (Fig. 4.5 E). Band 3 showed a peak a typical emission peak of PSI at 712 nm; however, the emission yield was greatly reduced relative to the major peak at 682 nm which is indicative of free LHCII. Thus, PSI from HS-acclimated C. reinhardtii appears to be functionally distinct from that of LS-acclimated cells, resembling previous reports on PSI 77K fluorescence in whole cells and isolated pigment complexes from UWO241. Fluorescence emission of the sucrose density bands isolated from both psychrophiles exhibited some novel features (Fig. 4.5 F,G). While Band 1 fluorescence emission was typical of LHCII, compared with C. reinhardtii, Band 2 (PSII) was shifted to shorter emission wavelengths in both psychrophiles. Furthermore, in ICE-MDV-HS, both bands 3 and 4 both exhibited a broad shoulder in fluorescence emission between 705 - 712 nm. However, in HS-grown UWO241, band 3 (PSI) and band 4 (supercomplex) exhibit minimal to no fluorescence emission at longer wavelengths.

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4.3.6 Proteome analysis of high salinity-associated supercomplexes

Despite variable levels of salinity tolerances across the strains, we were able to detect both the presence of PSI supercomplexes and high CEF rates in all three Chlamydomonas spp. under steady-state salinity stress. Previously, this phenotype has been generally associated on with short-term state 2 conditions. We wondered if the protein components of these newly discovered supercomplexes were also comparable with the highly studies state 2 supercomplex. We compared supercomplexes isolated from both C. reinhardtii and UWO241 after acclimation to high salinity (HS) stress with C. reinhardtii under state 2 conditions (SII), using shotgun proteomics (Fig. 4.6 A). The relative abundance of several proteins belonging to four major

thylakoid protein complexes (PSI, LHCI, LHCII, Cyt b6f) which have been previously associated with other supercomplexes were compared using normalized spectral abundance factor (NSAF) (Zybailov et al., 2006). PSI subunits were the most abundant protein constituents of supercomplexes in all the 3 conditions (Fig. 4.6 B). C. reinhardtii SII had the highest levels of the core reaction center protein, PsaA compared to the supercomplexes isolated from HS conditions. C. reinhardtii HS supercomplex displayed higher abundance of PsaC-N subunits compared to major PsaA and PsaB subunits. On the other hand, UWO241-HS supercomplex only contained 6 subunits out of the 11 PSI subunits and completely lacked PsaA and B. LHCI subunit abundance was also variable among the different supercomplexes. While C. reinhardtii - SII supercomplex contained all but one Lhca subunit, C. reinhardtii -HS only contained 4 and UWO241 only contained 2 Lhca subunits (Fig. 4.6 C). As predicted, C. reinhardtii -SII supercomplex contains the 4 major LHCII subunits implicated in supercomplex of state 2 (Fig. 4.6 D). Surprisingly, these Lhcb proteins were also detected in the C. reinhardtii -HS supercomplex. In contrast, the UWO241 supercomplex only contained the inner antenna protein cp29, which has the lowest abundance in UWO241 compared to both C. reinhardtii supercomplexes. Last, several cytb6f subunits were present in all three supercomplexes, however UWO241 had remarkedly higher abundance of the 3 subunits present in its supercomplex (PetA, B and C). Furthermore, PETO, which has been implicated to be important in supercomplex formation under anaerobiosis, was only present in the C. reinhardtii supercomplex. Our results show that the supercomplex from UWO241 is different from both C. reinhardtii supercomplexes; however, the HS supercomplexes of C. reinhardtii and UWO241 share some similarities in PSI and LHCI subunit composition.

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4.4 DISCUSSION

In this study, we employed three different Chlamydomonas species with variable salinity sensitivity to understand the role of CEF and supercomplex formation during acclimation to long-term stress. To acclimate the algae to high salinity conditions, we first conducted growth curves using a salinity gradient. We identified the maximum salinity tolerance as the highest NaCl concentration where the organism showed logarithmic growth. As known previously, for UWO241, maximum salt concentration with log growth is 700 mM NaCl (Fig. 4.1). Interestingly, the sister Chlamydomonas ICE-MDV tolerated salinity up to 500 mM NaCl but failed to grow at 700 mM NaCl. Thus ICE-MDV is halotolerant but more sensitive to high salt concentrations. We attribute the differential salinity sensitivities between the two Lake Bonney algae to the strict stratification of salinity and nutrient concentrations in the lake (Priscu & Spigel, 1996). ICE-MDV was isolated from, and dominates, the nutrient poor, freshwater environment of the shallow layers in Lake Bonney. On the other hand, UWO241 is found at deeper hypersaline layers of Lake Bonney (Cvetkovska, Smith, & Huner, 2017). Thus, the two lake Bonney algae have evolved to survive under different conditions which are specific stratified layers in the lake water column. State transitions are short-term acclimatory mechanisms that help photosynthetic organisms balance excitation energy in the presence of high light stress. Broadly diverse photosynthetic lifeforms have retained the ability to utilize this mechanism to survive short-term stress. Early studies showed that as a consequence of isolation under permanent conditions for perhaps thousands of years, UWO241 lacks the ability to undergo a classic state transition and exhibits a unique 77K fluorescence spectrum with a significantly attenuated PSI fluorescence. While ICE-MDV has evolved in the same lake and is also a psychrophile, unlike UWO241 ICE- MDV exhibited a typical 77K spectra with a prominent PSI fluorescence emission which was modulated under the state transition treatment (Fig. 4.3 B). Thus, despite long-term isolation under the ice cover of Lake Bonney, ICE-MDV has retained a normal state transition response. In addition, state transitions require reversible phosphorylation of LHCII proteins. In contrast with UWO241 phospho-LHCII proteins were detected in ICE-MDV thylakoids (Fig. 4.4). While UWO241 and ICE-MDV exhibited differential capacities for state transitions and altered PSI functional organization under low salinity; following acclimation to high salinity, ICE-MDV exhibited alterations in PSI functional organization which was comparable with

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UWO241, ie. loss of PSI fluorescence. Moreover, both 77K fluorescence emission and PSII room temperature fluorescence results indicated that the state transition response was also significantly attenuated (3.5-fold) in ICE-MDV in high salinity (Fig. 4.3 B, D). In their natural environment, ICE-MDV is exposed to higher PAR intensities in the shallow layers of lake Bonney, compared to UWO241 which is shade adapted in the deeper layers. In addition, light variation caused by ice thickness and/or transparency is likely more pronounced in the shallow layers of the water column. Thus, ICE-MDV has retained the ability for typical state transitions under variable light and freshwater conditions (ie. low salinity) of its natural habitat. However, the attenuation of this response after salinity acclimation suggests that there is a common effect of salinity on the organization of the photosynthetic apparatus in both halo-tolerant Antarctic algae. Interestingly, even HS-acclimated cultures of the salt-sensitive model green alga C. reinhardtii exhibited lower PSI peak after salinity acclimation compared to low salinity (Fig. 4.3 C) and a reduction in state transition capacity (Fig. 4.3 F). This decrease in state transition potential is not associated with damaged PSII as the FM under low and high salinity were not significantly different (Appendix Fig. 4.2). These changes in short-term stress response after long-term salt stress acclimation in three different Chlamydomonas species all suggest a common salinity-induced effect on reorganization of the photosynthetic apparatus which impairs the capacity for state transition. This lesion in state transitions is most pronounced in UWO241, the alga which possesses the highest tolerance to salinity stress. What changes in photosynthetic light reactions might be contributing to altered state transition response? A major consequence of a state transition response is formation of PSI- LHCI-LHCII supercomplex that helps increase CEF under high light conditions (Iwai et al., 2010). Higher CEF is then used to balance excitation energy between the two photosystems through initiation of non-photochemical quenching (NPQ) (Ebenhöh, Fucile, Finazzi, Rochaix, & Goldschmidt-Clermont, 2014). The pioneering research led by Takahashi et al., in 2013 elucidated that although both CEF and state transitions are controlled through redox status of the plastoquinone pool, they can occur independent of each other (Takahashi et al., 2013). This major study changed the course of CEF research and paved way for understanding CEF in the context of other major stresses. Recent studies have established that a restructured photosynthetic apparatus that is primed for constitutive capacity for CEF is key in high salinity acclimation in UWO241 (Kalra et al., 2020). Based on these results, we surmised that increased CEF could be

187 triggered in other Chlamydomonas species exposed to high salinity. We also suggested that stable PSI-supercomplexes are formed during steady-state high salinity conditions that help increase CEF. Restructuring of the photosynthetic apparatus to provide sustained CEF, may inhibit the state transition response. CEF rates were determined in all three Chlamydomonas species using P700 oxidation/reduction kinetics in presence of DCMU to inhibit LEF (Fig. 4.2). We showed that the psychrophiles have markedly fast CEF rates, compared to mesophile C. reinhardtii under control low salt conditions. Remarkably, CEF rates increased significantly under high salinity condition in all three species (Fig. 4.2). Thus, CEF may play a ubiquitous role in long-term salinity acclimation and is not a unique adaptation in the halotolerant UWO241. Recently, studies on tomato plants have also revealed cross-protective role of CEF in salinity and drought tolerance (Yang et al., 2020). The authors showed that increased CEF and NPQ capacity is maintained after salinity stress in tomato leaves, which further primes the plant to better tolerate drought stress. We suggest that increased CEF in high salinity acclimated photosynthetic organisms may help prime the organism for higher CEF rates, keeping the electron transport chain oxidized and therefore reducing the need for state transitions. As high salinity acclimation is linked with increased CEF rates in Chlamydomonas species, we wondered whether PSI-supercomplexes are also formed to support high rates of CEF. PSI-supercomplexes are extensively studied in C. reinhardtii under state 2 or anaerobic conditions. From the first biochemical characterization of PSI-supercomplex in state 2 condition by Iwai et al in 2010, to recent advances in structural characterization in anaerobic and state 2 condition, the last decade has accelerated our understanding of molecular basis for CEF in C. reinhardtii (Z. Huang et al., 2021; Iwai et al., 2010; Steinbeck et al., 2018; Takahashi et al., 2016; Terashima et al., 2012). Many components of the PETC were shown to be associated with

PSI-supercomplexes under anoxia and state 2: PSI, cytochrome b6f, LHCI, LHCII, CAS (Calcium Sensing protein), FNR1 (Ferredoxin NADP Reductase), ANR1 (Anaerobic response I protein). However, whether a similar supercomplex operates in conditions of long-term steady state stress is unknown. We addressed this question by probing thylakoid membranes isolated from HS-acclimated cells of Chlamydomonas species, all capable of HS-associated high CEF. Remarkably, a high molecular weight band that migrated lower than PSI-LHCI bands was detected in all three of the Chlamydomonas species (Fig. 4.5 A, B). The HS-associated

188 supercomplex from C. reinhardtii was distinct from the State 2 supercomplex: PSI fluorescence was greatly attenuated in both PSI and the supercomplex of under the HS-condition. These results matched that of whole cells, which also displayed loss of PSI fluorescence emission under HS (Fig. 4.3 C). On the other hand, although both high salinity acclimated ICE-MDV and UWO241 gradients contained all four bands including the supercomplex, they had similar banding patterns which distinct from C. reinhardtii, including denser LHCII that migrated lower in the gradients (Fig. 4.5 B). In addition, band 3 and 4 (ie. PSI and supercomplexes, respectively) also showed attenuated PSI fluorescence emission (Fig. 4.5 E, F). The long wavelength fluorescence emission peak at 712 nm at 77K corresponds to the PSI-LHCI complex in Chlamydomonas. Salinity stress has been shown to affect the PSI-LHCI complex through degradation of several PSI and LHCI subunits (Subramanyam et al., 2010). We have previously UWO241 whole cells and supercomplex fractions lack most Lhca proteins (Kalra et al., 2020). Selective loss of Lhca proteins might be responsible for attenuated fluorescence peak in high salinity acclimated Chlamydomonas species. In this study, we have shown for the first time that PSI-supercomplexes are associated with increased CEF in different Chlamydomonas species after acclimation to high salinity. Early research on identification of the state 2 supercomplex of C. reinhardtii revealed the presence of several LHCII subunits that migrate from PSII to PSI during state 2 transition (Takahashi, Iwai, Takahashi, & Minagawa, 2006). Later studies identified additional protein participants in the supercomplex. The state 2 supercomplex was shown to be calcium-regulated, containing CAS, ANR1 and PGRL1 proteins (Terashima et al. 2012). Further research discovered that cytochrome b6f is also essential for supercomplex formation (Minagawa, 2016). A recent structural study identified two LHCI (Lhca2 and 9) subunits whose dissociation is important for PSI-LHCI-cytb6f supercomplex formation (Steinbeck et al., 2018). In addition, a PSI-LHCI-LHCII supercomplex of C. reinhardtii under state 2 was shown to contain two LHCII trimers and ten LHCI subunits (Z. Huang et al., 2021). To understand whether the high salt supercomplexes share similarities with the state 2 supercomplex, we used shotgun proteomics to identify the supercomplex components. The band 4 supercomplex fractions of C. reinhardtii and UWO241 high salt (HS) sucrose gradients were compared to state 2 (SII) supercomplex of C. reinhardtii (Fig. 4.6 A). Overall, C. reinhardtii SII and HS supercomplex fractions shared many similarities in their protein composition, however UWO241 HS supercomplex was somewhat different (Table 1).

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PSI and LHCI form the core of the supercomplex assembly in all CEF supercomplexes characterized (Z. Huang et al., 2021; Iwai et al., 2010; Steinbeck et al., 2018; Terashima et al., 2012). C. reinhardtii has a significantly larger PSI-LHCI, with many additional Lhca proteins, relative to higher plants. Both HS and SII C. reinhardtii supercomplexes contained 10 out of 13 PSI subunits, whereas UWO241 supercomplex only contained 4. On the other hand, LHCI subunits were significantly reduced in HS supercomplexes compared to SII supercomplex of C. reinhardtii. Many photosynthetic organisms exhibit different pigment-protein composition and antenna sizes of PSI-LHCI. Here we show that salinity-dependent reduction in Lhca’s are likely to contribute to reduced PSI peak in HS supercomplex 77K fluorescence emission spectra.

Recently, cytochrome b6f has been shown to be an important member of the state 2 supercomplex, where electron transfer activity revealed reduction of cytb subunit through ferredoxin (Minagawa, 2016). Both HS supercomplexes contained several cytochrome b6f subunits; however, UWO241 supercomplex had disproportionately higher abundance of Pet A, B and C subunits, and only trace levels of the other Pet proteins (Table 1, Fig. 4.6 E). During state transitions, LHCII proteins migrate from PSII to PSI and thus form crucial components of state 2 supercomplex. Interestingly even HS supercomplex of C. reinhardtii contained several LHCII subunits that were in high abundance (Table 1, Fig. 4.6 D). FNR1, CAS and ANR1 proteins have been shown to co-fractionate with the state 2 supercomplex (Terashima et al., 2012). The HS supercomplex of C. reinhardtii contained all three of these proteins, suggesting involvement of similar supercomplex proteins across treatments. Previous research on state 2 supercomplex showed CAS protein to be involved in CEF initiation, and its absence inhibited CEF response (Terashima et al., 2012). Moreover, CAS was shown to interact with ANR1 protein and CEF was regulated in a Ca2+-dependent manner. Presence of CAS and ANR1 in the HS supercomplex suggests a possible interaction between these proteins. Interestingly, CAS and Ca2+ were also shown to be important in high light acclimation and in regulation of LHCSR3 (Light Harvesting Complex Stress Response 3) protein involved in NPQ response (Petroutsos et al., 2011). More recently, LHCSR3 protein was shown to be involved in both PSII and PSI supercomplexes (Girolomoni et al., 2019). Surprisingly, LHCSR3 protein is also found in the HS supercomplex of C. reinhardtii (Table 1). This protein has been also shown to be important in high salinity stress adaptation in Chlamydomonas sp. ICE-L (Mou et al., 2012). Furthermore, Ca2+ is also involved as a cofactor in oxygen evolving complex (OEC) of PSII,

190 especially the OEC subunits PSBP and PSBO are involved in Ca2+ binding. The PSBP protein was also previously shown to be involved in UWO241 HS supercomplex but its role in supercomplex formation is still unknown. We found that PSBP and PSBQ were present in both HS and SII supercomplex of C. reinhardtii (Table 1). We suggest that calcium signaling, and homeostasis through CAS, ANR1, LHCSR3 involvement, also plays a role in supercomplex formation during long-term salinity stress. Further research should be conducted to understand whether calcium homeostasis through CAS is essential for CEF regulation under long-term salinity stress. In this study we have shown that CEF is important in long-term salinity acclimation in three different Chlamydomonas species from various natural habitats, pointing towards ubiquity of CEF in salinity acclimation in green algae. We also showed that PSI-supercomplexes are associated with high salt acclimated cultures. Short-term stress responses like state transitions were also attenuated when cultures were acclimated to salinity, possibly as a result of restricted photosynthetic apparatus primed for CEF through PSI-supercomplexes. Lastly, we characterized the high salt supercomplex in C. reinhardtii, and showed that it shares many components with the state 2 supercomplex, including proteins involved in calcium homeostasis. We thus predict

PSI-LHCI-LHCII-cytb6f supercomplexes may regulate CEF in a variety of stress conditions, including short- and long-term, through a calcium regulated manner.

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UWO241 ICE-MDV C.reinhardtii 1.5 1.5 B 1.5 C BBM A BBM BBM 50 mM NaCl 250 mM NaCl 250 mM NaCl 100 mM NaCl 500 mM NaCl 500 mM NaCl 1.0 1.0 1.0 150 mM NaCl 700 mM NaCl 700 mM NaCl 200 mM NaCl

0.5 0.5 0.5 O.D. (750 nm) O.D. (750 nm) O.D. (750 nm)

0.0 0.0 0.0 0 5 10 15 0 5 10 15 0 1 2 3 4 5 6 7 Time (days) Time (days) Time (days)

Figure 4.1. Growth Curves under salinity gradient for the three Chlamydomonas species to identify maximum salinity tolerance. A. UWO241. B. ICE-MDV. C. C. reinhardtii. Growth was measured as optical density at 750 nm. BBM = 0.46 mM NaCl. (n=3, ±SE)

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A UWO241 B ICE-MDV C C. reinhardtii

600 600 600 **** **** ****

400 400 400 (ms) (ms) (ms) 1/2 1/2 t 1/2 t t 200 200 200

0 0 0

Low salt High salt Low salt High salt Low salt High salt D E F 0.8 0.8 *** 0.8 ns ****

0.6 0.6 0.6 820 820 820 0.4 0.4

0.4 A A A Δ Δ Δ

0.2 0.2 0.2

0.0 0.0 0.0

Low salt High salt Low salt High salt Low salt High salt

Figure 4.2. P700 oxidation/reduction analysis on the three Chlamydomonas spp. under low and high salinity. Top panel: Re-reduction rate was calculated under low (black) and high salinity (grey) for all three strains: UWO241 (A), ICE-MDV (B), C. reinhardtii (C). Bottom panel: The proportion of photooxidizable P700 is shown as change in absorbance at 820 nm for all three strains under low and high salinity: UWO241 (D), ICE-MDV (E), C. reinhardtii (F). Actinic red light was used with DCMU to inhibit electron flow from PSII. (n=9, ± SD, ns (not significant, p > 0.05), ** (p<0.01), *** (p<0.005), **** (p<0.001))

193

UWO241 ICE-MDV C. reinhardtii

1.2 1.2 1.2 A Low salt State I B C 1.0 Low salt State II 1.0 1.0 0.8 High salt State I 0.8 0.8 High salt State II 0.6 0.6 0.6

0.4 0.4 0.4

0.2 0.2 0.2 Fluorescence (R.F.U) Fluorescence (R.F.U) Fluorescence (R.F.U) 0.0 0.0 0.0 660 680 700 720 740 760 660 680 700 720 740 760 660 680 700 720 740 760 Wavelength (nm) Wavelength (nm) Wavelength (nm) D ✱✱✱ E 50 F 0 50 %

% ✱✱

% 40 ST1 ST1 -10 40 M M ST1 M

)/ F 30 )/ F ns 30 )/ F ST2

ST2 -20 M M ST2 20 M 20 - F - F

-30 - F ST1 ST1 10 10 M M ST1 M (F (F -40 (F 0 0

Low salt Low salt High salt Low salt High salt High salt

Figure 4.3. State transition tests after acclimation to low and high salinity in Chlamydomonas species. Top panel: Low temperature (77K) fluorescence spectra of the three Chlamydomonas spp. under state I and state II conditions after low and high salinity acclimation. Fluorescence values are shown as relative fluorescence units (R.F.U) for each strain: UWO241 (A); ICE-MDV (B); C.reinhardtii (C). Low salinity - Black, High salinity - Red. State I – Closed line, State II – dotted line. Bottom panel: Maximal capacity for switching LHCII antenna during State transition induction calculated using room temperature PSII maximum

fluorescence (FM) as described before (ref) for each strain: UWO241 (D); ICE-MDV (E); C. reinhardtii (F). ST1: state 1, ST2: state 2. (n=4-6; ±SD; ns (not significant, p > 0.05), ** (p<0.01), *** (p<0.005))

194 kDa LS HS LS HS LS HS 250 150

100 75

37 CP 26 CP 29 Type II Type I 25 Type IV Type III 15

UWO241 ICE-MDV C. reinhardtii

Figure 4.4. Thylakoid phosphorylation pattern of the three Chlamydomonas spp. under low (LS) and high (HS) salinity. Isolated thylakoids were run on 12% SDS-PAGE and probed with phospho-threonine antibody. Mol. wt ladder (KDa) is shown on the left. The different LHCII types are labelled on the right.

195

A LS-SI LS -SII HS B HS HS

1 1 1 2 2 2 3 3 3

4 4

C. reinhardtii ICE-MDVUWO241

ICE-MDV UWO241 C. reinhardtii C. reinhardtii C. reinhardtii HS HS LS-SI LS-SII HS 1.2 1.2 1.2 1.2 1.2 F G C D Band 1 E Band 1 Band 1 Band 1 Band 1 1.0 1.0 1.0 1.0 Band 2 1.0 Band 2 Band 2 Band 2 Band 2 0.8 0.8 0.8 0.8 Band 3 0.8 Band 3 Band 3 Band 3 Band 3 Band 4 0.6 0.6 Band 4 0.6 0.6 0.6 Band 4 Band 4 0.4 0.4 0.4 0.4 0.4 0.2

0.2 Fluorescence (R.F.U) 0.2 0.2 0.2 Fluorescence (R.F.U) Fluorescence (R.F.U) Fluorescence (R.F.U) Fluorescence (R.F.U) 0.0 0.0 0.0 0.0 0.0 660 680 700 720 740 760 660 680 700 720 740 760 660 680 700 720 740 760 660 680 700 720 740 760 660 680 700 720 740 760 Wavelength (nm) Wavelength (nm) Wavelength (nm) Wavelength (nm) Wavelength (nm)

Figure 4.5. Isolation of supercomplexes from conditions promoting CEF in Chlamydomonas species. Top panel: Separation of protein complexes on a sucrose density gradient for A. model mesophile C. reinhardtii and B. the psychrophiles C. sp. ICE-MDV and C. sp. UWO241. Bottom panel: 77K fluorescence spectra for protein complex bands isolated from sucrose density gradient: C. reinhardtii under low salt and state 1 (A), state 2 (B), under high salinity (E); ICE-MDV under high salinity (F); UWO241 under high salinity (G). LS-SI: Low salinity, state 1; LS-SII: Low salinity, state 2; HS: High salinity. Band 1: LHCII, Band 2: PSII, Band 3: PSI-LHCI, Band 4: Supercomplex.

196

0.06 UWO-HS 0.16 B C UWO-HS C.r-HS C.r-HS 0.05 C.r-SII C.r-SII

0.12 A 0.04

0.03 0.08

C. r -SII C. r-HS UWO-HS NSAF NSAF 0.02 0.04

0.01

0.00 ABCDEFGHKLN 0.00 Lhca1 Lhca2 Lhca3 Lhca4 Lhca5 Lhca7 Lhca9 PSI Subunits LHCI subunits 0.06 0.06 D UWO-HS E UWO-HS C.r-HS C.r-HS 0.05 0.05 C.r-SII C.r-SII Supercomplex 0.04 0.04

0.03 0.03 NSAF NSAF 0.02 0.02

0.01 0.01

0.00 0.00 Lhcbm3 Lhcbm5 Cp26 Cp29 PetA PetB PetC PetD PetM PetO LHCII Cytb f subunits 6

Figure 4.6. Proteome comparison of supercomplex fractions from C. reinhardtii-state 2, C. reinhardtii- HS and UWO241-HS. A. Sucrose density gradients highlighting the supercomplex fraction analyzed. The normalized spectral abundance factor (NSAF) for each identified protein within a supercomplex were calculated. The major photosynthetic protein-complexes and their subunits participating in supercomplex formation are shown: Photosystem I (PSI, B), Light Harvesting complex I (LHCI, C), Light Harvesting

complex II (LHCII, D) and Cytochrome b6f (Cytb6f, E). UWO241-HS (UWO-HS, Black), C. reinhardtii-HS (C.r-HS, White), C.reinhardtii state 2 (C.r-SII, Grey).

197

TABLE 4.1. Major proteins involved in supercomplex formation in C. reinhardtii and

UWO241. The subunits of each protein identified through shot-gun proteomics are shown. HS: High salinity acclimated; State 2: State 2 locked culture; PSI: photosystem I; LHCI: Light harvesting complex I; LHCII: Light harvesting complex II, Cyt b6f: cytochrome b6f, FNR: Ferredoxin NADP reductase, FtsH: ATP dependent zinc metalloprotease, OEEP: Oxygen evolving enhancer protein, CAS: Calcium sensing protein

Protein complex UWO241-HS C. reinhardtii -HS C. reinhardtii -State 2

PSI PsaD, PsaF, PsaE, PsaK, PsaA, PsaB, PsaC, PsaA, PsaB, PsaC, PsaH PsaD, PsaE, PsaF, PsaD, PsaE, PsaF, PsaG, PsaK, PsaL, PsaG, PsaK, PsaL, PsaN PsaN LHCI Lhca3, Lhca5 Lhca1, Lhca2, Lhca7, Lhca1, Lhca2, Lhca3, Lhca9 Lhca4, Lhca5, Lhca7, Lhca9 LHCII Cp29 Lhcbm1, Lhcbm3, Lhcbm3, Lhcbm5, Lhcbm5, CP26, CP29, CP26, CP29 Lhcsr3

Cyt b6f Pet A, PetB, PetC PetA, PetB, PetC, PetA, PetB, PetC, PetD, PetM, PetO PetD, PetM, PetO FNR * FNR1 FNR1 FtsH FtsH2, FtsH 5 FtsH1, 2 FtsH1, 2, 4 OEEP PsbP, PsbQ PsbQ, PsbP1 PsbQ, PsbO, PsbP1 Calcium sensing CAS CAS * receptor

198

4.5 REFERENCES

Aboelmy, M. H., & Peterhansel, C. (2014). Enzymatic characterization of Chlamydomonas reinhardtii glycolate dehydrogenase and its nearest proteobacterial homologue. Plant Physiology and Biochemistry, 79, 25–30. https://doi.org/10.1016/j.plaphy.2014.03.009 Aguilera, A., Souza-Egipsy, V., & Amils, R. (2012). Photosynthesis in Extreme Environments. Artificial Photosynthesis. https://doi.org/10.5772/26940 Airaki, M., Leterrier, M., Mateos, R. M., Valderrama, R., Chaki, M., Barroso, J. B., … Corpas, F. J. (2012). Metabolism of reactive oxygen species and reactive nitrogen species in pepper (Capsicum annuum L.) plants under low temperature stress. Plant, Cell and Environment, 35(2), 281–295. https://doi.org/10.1111/j.1365-3040.2011.02310.x Alexandratos, N., & Brunismas, J. (2012). WORLD AGRICULTURE TOWARDS 2030/2050. JESA Working Papers 12-03. https://doi.org/10.22004/ag.econ.288998 Allakhverdiev, S. I., & Murata, N. (2008). Salt stress inhibits photosystems II and I in cyanobacteria. Photosynthesis Research, 98(1–3), 529–539. https://doi.org/10.1007/s11120- 008-9334-x Alric, J. (2010). Cyclic electron flow around photosystem I in unicellular green algae. Photosynthesis Research, 106(1–2), 47–56. https://doi.org/10.1007/s11120-010-9566-4 Alric, J. (2015). The plastoquinone pool, poised for cyclic electron flow? Frontiers in Plant Science, 6(JULY), 1–4. https://doi.org/10.3389/fpls.2015.00540 Alric, J., Lavergne, J., & Rappaport, F. (2010). Redox and ATP control of photosynthetic cyclic electron flow in Chlamydomonas reinhardtii (I) aerobic conditions. Biochimica et Biophysica Acta - Bioenergetics, 1797(1), 44–51. https://doi.org/10.1016/j.bbabio.2009.07.009 Bai, X., Yang, L., Yang, Y., Ahmad, P., Yang, Y., & Hu, X. (2011). Deciphering the protective role of nitric oxide against salt stress at the physiological and proteomic levels in maize. Journal of Proteome Research, 10(10), 4349–4364. https://doi.org/10.1021/pr200333f Bonente, G., Pippa, S., Castellano, S., Bassi, R., & Ballottari, M. (2012). Acclimation of Chlamydomonas reinhardtii to different growth irradiances. Journal of Biological Chemistry, 287(8), 5833–5847. https://doi.org/10.1074/jbc.M111.304279 Cardona, T., Shao, S., & Nixon, P. J. (2018). Enhancing photosynthesis in plants: the light reactions. Essays in Biochemistry, 62(1), 85–94. https://doi.org/10.1042/EBC20170015

199

Carvalho, P. C., Lima, D. B., Leprevost, F. V., Santos, M. D. M., Fischer, J. S. G., Aquino, P. F., … Barbosa, V. C. (2016). Integrated analysis of shotgun proteomic data with PatternLab for proteomics 4.0. Nature Protocols, 11(1), 102–117. https://doi.org/10.1038/nprot.2015.133 Chang, R. L., Ghamsari, L., Manichaikul, A., Hom, E. F. Y., Balaji, S., Fu, W., … Papin, J. A. (2011). Metabolic network reconstruction of Chlamydomonas offers insight into light-driven algal metabolism. Molecular Systems Biology, 7(518). https://doi.org/10.1038/msb.2011.52 Chen, H., Hu, J., Qiao, Y., Chen, W., Rong, J., Zhang, Y., … Wang, Q. (2015). Ca 2+ -regulated cyclic electron flow supplies ATP for nitrogen starvation-induced lipid biosynthesis in green alga. Scientific Reports, 5(October), 1–12. https://doi.org/10.1038/srep15117 Chen, X., Tian, D., Kong, X., Chen, Q., Abd, A. A., Hu, X., & Jia, A. (2016). The role of nitric oxide signalling in response to salt stress in Chlamydomonas reinhardtii. Planta, 244(3), 651– 669. https://doi.org/10.1007/s00425-016-2528-0 Cook, G., Teufel, A., Kalra, I., Li, W., Wang, X., Priscu, J., & Morgan-Kiss, R. (2019). The Antarctic psychrophiles Chlamydomonas spp. UWO241 and ICE-MDV exhibit differential restructuring of photosystem I in response to iron. Photosynthesis Research, 141(2), 209– 228. https://doi.org/10.1007/s11120-019-00621-0 Cvetkovska, M., Smith, R., & Huner, N. P. A. (2017). Chilling out : the evolution and diversification of psychrophilic algae with a focus on Chlamydomonadales, 1169–1184. https://doi.org/10.1007/s00300-016-2045-4 Dietzel, L., Bräutigam, K., & Pfannschmidt, T. (2008). Photosynthetic acclimation: State transitions and adjustment of photosystem stoichiometry - Functional relationships between short-term and long-term light quality acclimation in plants. FEBS Journal, 275(6), 1080– 1088. https://doi.org/10.1111/j.1742-4658.2008.06264.x Dijkhuizen, L., & Harder, W. (1985). Current views on the regulation of autotrophic carbon dioxide fixation via the Calvin cycle in bacteria. Antonie van Leeuwenhoek, 50(1984), 473– 487. Dolhi, J. M., Maxwell, D. P., & Morgan-Kiss, R. M. (2013). Review: The Antarctic Chlamydomonas raudensis: An emerging model for cold adaptation of photosynthesis. Extremophiles, 17(5), 711–722. https://doi.org/10.1007/s00792-013-0571-3 Dolhi, J. M., Teufel, A. G., Kong, W., & Morgan-Kiss, R. M. (2015). Diversity and spatial

200

distribution of autotrophic communities within and between ice-covered Antarctic lakes (McMurdo Dry Valleys). Limnology and Oceanography, 60(3), 977–991. https://doi.org/10.1002/lno.10071 Ebenhöh, O., Fucile, G., Finazzi, G., Rochaix, J. D., & Goldschmidt-Clermont, M. (2014). Short- term acclimation of the photosynthetic electron transfer chain to changing light: A mathematical model. Philosophical Transactions of the Royal Society B: Biological Sciences, 369(1640). https://doi.org/10.1098/rstb.2013.0223 Ensminger, I., Busch, F., & Huner, N. P. A. (2006). Photostasis and cold acclimation: Sensing low temperature through photosynthesis. Physiologia Plantarum, 126(1), 28–44. https://doi.org/10.1111/j.1399-3054.2006.00627.x Erickson, E., Wakao, S., & Niyogi, K. K. (2015). Light stress and photoprotection in Chlamydomonas reinhardtii. Plant Journal, 82(3), 449–465. https://doi.org/10.1111/tpj.12825 Fang, W., Si, Y., Douglass, S., Casero, D., Merchant, S. S., Pellegrini, M., … Spaldinga, M. H. (2012). Transcriptome-wide changes in Chlamydomonas reinhardtii gene expression regulated by carbon dioxide and the CO2-concentrating mechanism regulator CIA5/CCM1. Plant Cell, 24(5), 1876–1893. https://doi.org/10.1105/tpc.112.097949 Finazzi, G., & Johnson, G. N. (2016). Cyclic electron flow: facts and hypotheses. Photosynthesis Research, 129(3), 227–230. https://doi.org/10.1007/s11120-016-0306-2 Fritsen, C. H., & Priscu, J. C. (1999). Seasonal change in the optical properties of the permanent ice cover on Lake Bonney, Antarctica: Consequences for lake productivity and phytoplankton dynamics. Limnology and Oceanography, 44(2), 447–454. https://doi.org/10.4319/lo.1999.44.2.0447 Girolomoni, L., Cazzaniga, S., Pinnola, A., Perozeni, F., Ballottari, M., & Bassi, R. (2019). LHCSR3 is a nonphotochemical quencher of both photosystems in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America, 116(10), 4212–4217. https://doi.org/10.1073/pnas.1809812116 Girolomoni, L., Ferrante, P., Berteotti, S., Giuliano, G., Bassi, R., & Ballottari, M. (2017). The function of LHCBM4/6/8 antenna proteins in Chlamydomonas reinhardtii. Journal of Experimental Botany, 68(3), 627–641. https://doi.org/10.1093/jxb/erw462

201

Godaux, D., Bailleul, B., Berne, N., & Cardol, P. (2015). Induction of photosynthetic carbon fixation in anoxia relies on hydrogenase activity and proton-gradient regulation-like1- mediated cyclic electron flow in Chlamydomonas reinhardtii. Plant Physiology, 168(2), 648– 658. https://doi.org/10.1104/pp.15.00105 Govindjee, Shevela, D., & Björn, L. O. (2017). Evolution of the Z-scheme of photosynthesis: a perspective. Photosynthesis Research, 133(1–3), 5–15. https://doi.org/10.1007/s11120-016- 0333-z Goyal, A. (2007a). Osmoregulation in Dunaliella, Part I: Effects of osmotic stress on photosynthesis, dark respiration and glycerol metabolism in Dunaliella tertiolecta and its salt- sensitive mutant (HL 25/8). Plant Physiology and Biochemistry, 45(9), 696–704. https://doi.org/10.1016/j.plaphy.2007.05.008 Goyal, A. (2007b). Osmoregulation in Dunaliella, Part II: Photosynthesis and starch contribute carbon for glycerol synthesis during a salt stress in Dunaliella tertiolecta. Plant Physiology and Biochemistry, 45(9), 705–710. https://doi.org/10.1016/j.plaphy.2007.05.009 Goyal, A., & Tolbert, N. E. (1990). Salicylhydroxamic acid (SHAM) inhibition of the dissolved inorganic carbon concentrating process in unicellular green algae. Plant Physiology, 92(3), 630–636. https://doi.org/10.1104/pp.92.3.630 Goyal, A., & Tolbert, N. E. (1996). Association of glycolate oxidation with photosynthetic electron transport in plant and algal chloroplasts. Proceedings of the National Academy of Sciences of the United States of America, 93(8), 3319–3324. https://doi.org/10.1073/pnas.93.8.3319 Gururani, M. A., Venkatesh, J., & Tran, L. S. P. (2015). Regulation of photosynthesis during abiotic stress-induced photoinhibition. Molecular Plant, 8(9), 1304–1320. https://doi.org/10.1016/j.molp.2015.05.005 He, Y., Fu, J., Yu, C., Wang, X., Jiang, Q., Hong, J., … Jiang, D. (2015). Increasing cyclic electron flow is related to Na+ sequestration into vacuoles for salt tolerance in soybean. Journal of Experimental Botany, 66(21), 6877–6889. https://doi.org/10.1093/jxb/erv392 Hema, R., Senthil-Kumar, M., Shivakumar, S., Chandrasekhara Reddy, P., & Udayakumar, M. (2007). Chlamydomonas reinhardtii, a model system for functional validation of abiotic stress responsive genes. Planta, 226(3), 655–670. https://doi.org/10.1007/s00425-007-0514-2 Ho, S. H., Nakanishi, A., Kato, Y., Yamasaki, H., Chang, J. S., Misawa, N., … Kondo, A. (2017).

202

Dynamic metabolic profiling together with transcription analysis reveals salinity-induced starch-To-lipid biosynthesis in alga Chlamydomonas sp. JSC4. Scientific Reports, 7(October 2016), 1–7. https://doi.org/10.1038/srep45471 Huang, W., Fu, P. L., Jiang, Y. J., Zhang, J. L., Zhang, S. B., Hu, H., & Cao, K. F. (2013). Differences in the responses of photosystem I and photosystem II of three tree species Cleistanthus sumatranus, Celtis philippensis and Pistacia weinmannifolia exposed to a prolonged drought in a tropical limestone forest. Tree Physiology, 33(2), 211–220. https://doi.org/10.1093/treephys/tps132 Huang, W., Zhang, S. B., & Cao, K. F. (2010). Stimulation of cyclic electron flow during recovery after chilling-induced photoinhibition of PSII. Plant and Cell Physiology, 51(11), 1922–1928. https://doi.org/10.1093/pcp/pcq144 Huang, W., Zhang, S. B., Xu, J. C., & Liu, T. (2017). Plasticity in roles of cyclic electron flow around photosystem I at contrasting temperatures in the chilling-sensitive plant Calotropis gigantea. Environmental and Experimental Botany, 141(May), 145–153. https://doi.org/10.1016/j.envexpbot.2017.07.011 Huang, Z., Liangliang, S., Wenda, W., Zhiyuan, M., Xiaohan, Y., Tingyun, K., … Guangye, H. (2021). Structure of photosystem I-LHCI-LHCII from the green alga Chlamydomonas reinhardtii in State 2. Nature Communications, (12(1)), 1–14. https://doi.org/10.1038/s41467- 021-21362-6 Hüner, N. P. A., Bode, R., Dahal, K., Hollis, L., Rosso, D., Krol, M., & Ivanov, A. G. (2012). Chloroplast redox imbalance governs phenotypic plasticity: The “grand design of photosynthesis” revisited. Frontiers in Plant Science, 3(NOV), 1–12. https://doi.org/10.3389/fpls.2012.00255 Huner, N. P. A., Öquist, G., & Sarhan, F. (1998). Energy balance and acclimation to light and cold, 1385(98). Ivanov, A. G., Morgan, R. M., Gray, G. R., Velitchkova, M. Y., & Huner, N. P. A. (1998). Temperature/light dependent development of selective resistance to photoinhibition of photosystem I. FEBS Letters, 430(3), 288–292. https://doi.org/10.1016/S0014- 5793(98)00681-4 Iwai, M., Takizawa, K., Tokutsu, R., Okamuro, A., Takahashi, Y., & Minagawa, J. (2010).

203

Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis. Nature, 464(7292), 1210–1213. https://doi.org/10.1038/nature08885 Jeffrey, S. W., & Humphrey, G. F. (1975). New spectrophotometric equations for determining chlorophylls a, b, c1 and c2 in higher plants, algae and natural phytoplankton. Biochemie Und Physiologie Der Pflanzen, 167(2), 191–194. https://doi.org/10.1016/s0015-3796(17)30778-3 Kalra, I., Wang, X., Cvetkovska, M., Jeong, J., McHargue, W., Zhang, R., … Morgan-Kissa, R. (2020). Chlamydomonas sp. UWO 241 exhibits high cyclic electron flow and rewired metabolism under high salinity. Plant Physiology, 183(2), 588–601. https://doi.org/10.1104/pp.19.01280 Kato, Y., Ho, S. H., Vavricka, C. J., Chang, J. S., Hasunuma, T., & Kondo, A. (2017). Evolutionary engineering of salt-resistant Chlamydomonas sp. strains reveals salinity stress-activated starch-to-lipid biosynthesis switching. Bioresource Technology, 245, 1484–1490. https://doi.org/10.1016/j.biortech.2017.06.035 Khona, D. K., Shirolikar, S. M., Gawde, K. K., Hom, E., Deodhar, M. A., & D’Souza, J. S. (2016). Characterization of salt stress-induced palmelloids in the green alga, Chlamydomonas reinhardtii. Algal Research, 16(July), 434–448. https://doi.org/10.1016/j.algal.2016.03.035 Knopf, J. A., & Shapira, M. (2005). Degradation of Rubisco SSU during oxidative stress triggers aggregation of Rubisco particles in Chlamydomonas reinhardtii. Planta, 222(5), 787–793. https://doi.org/10.1007/s00425-005-0023-0 Kono, M., & Terashima, I. (2014). Long-term and short-term responses of the photosynthetic electron transport to fluctuating light. Journal of Photochemistry and Photobiology B: Biology, 137, 89–99. https://doi.org/10.1016/j.jphotobiol.2014.02.016 Kramer, D. M., & Evans, J. R. (2011). The importance of energy balance in improving photosynthetic productivity. Plant Physiology, 155(1), 70–78. https://doi.org/10.1104/pp.110.166652 Kumar, V., Wani, S. H., Penna, S., & Tran, L. S. P. (2018). Salinity Responses and Tolerance in Plants, Volume 1: Targeting Sensory, Transport and Signaling Mechanisms. Salinity Responses and Tolerance in Plants, Volume 1: Targeting Sensory, Transport and Signaling Mechanisms. https://doi.org/10.1007/978-3-319-75671-4 Kuo, E. Y. H., Chang, H. L., Lin, S. T., & Lee, T. M. (2020). High Light-Induced Nitric Oxide

204

Production Induces Autophagy and Cell Death in Chlamydomonas reinhardtii. Frontiers in Plant Science, 11(June), 1–21. https://doi.org/10.3389/fpls.2020.00772 Laemmli, U. K. (1970). Cleavage of Structural Proteins during the Assembly of the Head of Bacteriophage T4. Nature, 227, 680–685. Lemeille, S., & Rochaix, J. D. (2010). State transitions at the crossroad of thylakoid signalling pathways. Photosynthesis Research, 106(1–2), 33–46. https://doi.org/10.1007/s11120-010- 9538-8 León, R., & Galván, F. (1994). Halotolerance studies on Chlamydomonas reinhardtii: glycerol excretion by free and immobilized cells. Journal of Applied Phycology, 6(1), 13–20. https://doi.org/10.1007/BF02185898 Li, J., Tietz, S., Cruz, J. A., Strand, D. D., Xu, Y., Chen, J., … Hu, J. (2019). Photometric screens identified Arabidopsis peroxisome proteins that impact photosynthesis under dynamic light conditions. Plant Journal, 97(3), 460–474. https://doi.org/10.1111/tpj.14134 Li, W., & Morgan-Kiss, R. M. (2019). Influence of environmental drivers and potential interactions on the distribution of microbial communities from three permanently stratified Antarctic lakes. Frontiers in Microbiology, 10(MAY), 1–16. https://doi.org/10.3389/fmicb.2019.01067 Li, W., Podar, M., & Morgan-Kiss, R. M. (2016). Ultrastructural and single-cell-level characterization reveals metabolic versatility in a microbial eukaryote community from an ice-covered Antarctic lake. Applied and Environmental Microbiology, 82(12), 3659–3670. https://doi.org/10.1128/AEM.00478-16 Lima Neto, M. C., Cerqueira, J. V. A., da Cunha, J. R., Ribeiro, R. V., & Silveira, J. A. G. (2017). Cyclic electron flow, NPQ and photorespiration are crucial for the establishment of young plants of Ricinus communis and Jatropha curcas exposed to drought. Plant Biology, 19(4), 650–659. https://doi.org/10.1111/plb.12573 Liska, A. J., Shevchenko, A., Pick, U., & Katz, A. (2004). Enhanced photosynthesis and redox energy production contribute to salinity tolerance in Dunaliella as revealed by homology- based proteomics. Plant Physiology, 136(1), 2806–2817. https://doi.org/10.1104/pp.104.039438 Lizottel, M. P., & Prism, J. C. (1994). Natural fluorescence and quantum yields in vertically

205

stationary phytoplankton from perennially ice‐covered lakes. Limnology and Oceanography, 39(6), 1399–1410. https://doi.org/10.4319/lo.1994.39.6.1399 Lucker, B., & Kramer, D. M. (2013). Regulation of cyclic electron flow in Chlamydomonas reinhardtii under fluctuating carbon availability. Photosynthesis Research, 117(1–3), 449– 459. https://doi.org/10.1007/s11120-013-9932-0 Magneschi, L., Catalanotti, C., Subramanian, V., Dubini, A., Yang, W., Mus, F., … Grossman, A. R. (2012). A mutant in the ADH1 gene of Chlamydomonas reinhardtii elicits metabolic restructuring during anaerobiosis. Plant Physiology, 158(3), 1293–1305. https://doi.org/10.1104/pp.111.191569 Melis, A. (1991). Dynamics of photosynthetic membrane composition and function. BBA - Bioenergetics, 1058(2), 87–106. https://doi.org/10.1016/S0005-2728(05)80225-7 Minagawa, J. (2016). A Supercomplex of Cytochrome bf and Photosystem I for Cyclic Electron Flow, 453–462. https://doi.org/10.1007/978-94-017-7481-9_23 Morgan-Kiss, R., Ivanov, A. G., Williams, J., Khan, M., & Huner, N. P. A. (2002). Differential thermal effects on the energy distribution between photosystem II and photosystem I in thylakoid membranes of a psychrophilic and a mesophilic alga. Biochimica et Biophysica Acta - Biomembranes, 1561(2), 251–265. https://doi.org/10.1016/S0005-2736(02)00352-8 Morgan-Kiss, R. M., Ivanov, A. G., & Huner, N. P. A. (2002). The Antarctic psychrophile, Chlamydomonas subcaudata, is deficient in State I-State II transitions. Planta, 214(3), 435– 445. https://doi.org/10.1007/s004250100635 Morgan-Kiss, R. M., Ivanov, A. G., Modla, S., Czymmek, K., Hüner, N. P. A., Priscu, J. C., … Hanson, T. E. (2008). Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp., strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles, 12(5), 701–711. https://doi.org/10.1007/s00792-008-0176-4 Morgan-Kiss, R. M., Priscu, J. C., Pocock, T., Gudynaite-Savitch, L., & Huner, N. P. A. (2006). Adaptation and Acclimation of Photosynthetic Microorganisms to Permanently Cold Environments. Microbiology and Molecular Biology Reviews, 70(1), 222–252. https://doi.org/10.1128/mmbr.70.1.222-252.2006 Morgan, R. M., Ivanov, A. G., Priscu, J. C., Maxwell, D. P., & Huner, N. P. A. (1998). Structure and composition of the photochemical apparatus of the Antarctic green alga, Chlamydomonas

206

subcaudata. Photosynthesis Research, 56(3), 303–314. https://doi.org/10.1023/A:1006048519302 Morton, M. J. L., Awlia, M., Al-Tamimi, N., Saade, S., Pailles, Y., Negrão, S., & Tester, M. (2019). Salt stress under the scalpel – dissecting the genetics of salt tolerance. Plant Journal, 97(1), 148–163. https://doi.org/10.1111/tpj.14189 Mou, S., Zhang, X., Ye, N., Dong, M., Liang, C., Liang, Q., … Zheng, Z. (2012). Cloning and expression analysis of two different LhcSR genes involved in stress adaptation in an Antarctic microalga, Chlamydomonas sp. ICE-L. Extremophiles, 16(2), 193–203. https://doi.org/10.1007/s00792-011-0419-7 Müller, P., Li, X. P., & Niyogi, K. K. (2001). Non-photochemical quenching. A response to excess light energy. Plant Physiology, 125(4), 1558–1566. https://doi.org/10.1104/pp.125.4.1558 Munekage, Y., Hojo, M., Meurer, J., Endo, T., Tasaka, M., & Shikanai, T. (2002). PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in Arabidopsis. Cell, 110(3), 361–371. https://doi.org/10.1016/S0092-8674(02)00867-X Nawrocki, W. J., Liu, X., & Croce, R. (2020). Chlamydomonas reinhardtii exhibits de facto constitutive NPQ capacity in physiologically relevant conditions. Plant Physiology, 182(1), 472–479. https://doi.org/10.1104/pp.19.00658 Neale, P. J., & Melis, A. (1989). Salinity-stress Enhances Photoinhibition of Photosynthesis in Chlamydomonas reinhardtii. Journal of Plant Physiology, 134(5), 619–622. https://doi.org/10.1016/S0176-1617(89)80158-0 Neelam, S., & Subramanyam, R. (2013). Alteration of photochemistry and protein degradation of photosystem II from Chlamydomonas reinhardtii under high salt grown cells. Journal of Photochemistry and Photobiology B: Biology, 124, 63–70. https://doi.org/10.1016/j.jphotobiol.2013.04.007 Negrão, S., Schmöckel, S. M., & Tester, M. (2017). Evaluating physiological responses of plants to salinity stress. Annals of Botany, 119(1), 1–11. https://doi.org/10.1093/aob/mcw191 Oren, A. (2014). The ecology of Dunaliella in high-salt environments. Journal of Biological Research (Greece), 21(1), 1–8. https://doi.org/10.1186/s40709-014-0023-y Perrineau, M. M., Zelzion, E., Gross, J., Price, D. C., Boyd, J., & Bhattacharya, D. (2014). Evolution of salt tolerance in a laboratory reared population of Chlamydomonas reinhardtii.

207

Environmental Microbiology, 16(6), 1755–1766. https://doi.org/10.1111/1462-2920.12372 Petroutsos, D., Busch, A., Janßen, I., Trompelt, K., Bergner, S. V., Weinl, S., … Hippler, M. (2011). The chloroplast calcium sensor CAS is required for photoacclimation in Chlamydomonas reinhardtii. Plant Cell, 23(8), 2950–2963. https://doi.org/10.1105/tpc.111.087973 Petroutsos, D., Tokutsu, R., Maruyama, S., Flori, S., Greiner, A., Magneschi, L., … Minagawa, J. (2016). A blue-light photoreceptor mediates the feedback regulation of photosynthesis. Nature, 537(7621), 563–566. https://doi.org/10.1038/nature19358 Plouviez, M., Wheeler, D., Shilton, A., Packer, M. A., McLenachan, P. A., Sanz-Luque, E., … Guieysse, B. (2017). The biosynthesis of nitrous oxide in the green alga Chlamydomonas reinhardtii. Plant Journal, 91(1), 45–56. https://doi.org/10.1111/tpj.13544 Priscu, J. C. (1998). Ecosystem Dynamics in a Polar Desert, 72. Priscu, J. C., & Spigel, R. H. (1996). Evolution of temperature and salt structure of Lake Bonney, a chemically stratified Antarctic lake. Hydrobiologia, 321(3), 177–190. Priscu, J. C., Wolf, F., Takacs, D., & Fritsen, H. (1999). Carbon Transformatiin a Perennially Ice- Covered Antarctic Lake, 49(12), 997–1008. Raymond, J. A., & Morgan-Kiss, R. (2013). Separate Origins of Ice-Binding Proteins in Antarctic Chlamydomonas Species. PLoS ONE, 8(3), 6–11. https://doi.org/10.1371/journal.pone.0059186 Raymond, J. A., Morgan-Kiss, R., & Stahl-Rommel, S. (2020). Glycerol Is an Osmoprotectant in Two Antarctic Chlamydomonas Species From an Ice-Covered Saline Lake and Is Synthesized by an Unusual Bidomain Enzyme. Frontiers in Plant Science, 11(August), 1–8. https://doi.org/10.3389/fpls.2020.01259 Reynoso, G. T., & de Gamboa, B. A. (1982). Salt tolerance in the freshwater algae Chlamydomon as reinhardii: Effect of proline and taurine. Comparative Biochemistry and Physiology -- Part A: Physiology, 73(1), 95–99. https://doi.org/10.1016/0300-9629(82)90098-6 Roach, T., & Krieger-Liszkay, A. (2014). Regulation of Photosynthetic Electron Transport and Photoinhibition. Current Protein & Peptide Science, 15(4), 351–362. https://doi.org/10.2174/1389203715666140327105143 Rochaix, J. D. (2007). Role of thylakoid protein kinases in photosynthetic acclimation. FEBS

208

Letters, 581(15), 2768–2775. https://doi.org/10.1016/j.febslet.2007.04.038 Sabeeha S. Merchant1,*, Simon E. Prochnik2,*, Olivier Vallon3, Elizabeth H. Harris4, Steven A. Sanderfoot11, Martin H. Spalding12, Vladimir V. Kapitonov13, Qinghu Ren14, Patrick Laurence Maréchal-Drouard7, Wallace F. Marshall8, Liang-Hu Qu9, David R. Nels, A. (2007). The Chlamydomonas Genome Reveals the Evolution of Key. National Institutes of Health, 318(5848), 245–250. Saroussi, S. I., Wittkopp, T. M., & Grossman, A. R. (2016). The Type II NADPH dehydrogenase facilitates cyclic electron flow, energy-dependent quenching, and chlororespiratory metabolism during acclimation of Chlamydomonas reinhardtii to nitrogen deprivation. Plant Physiology, 170(4), 1975–1988. https://doi.org/10.1104/pp.15.02014 Shetty, P., Gitau, M. M., & Maróti, G. (2019). Salinity Stress Responses and Adaptation Mechanisms in Eukaryotic Green Microalgae. Cells, 8(12), 1–16. https://doi.org/10.3390/cells8121657 Shoun, H., Fushinobu, S., Jiang, L., Kim, S. W., & Wakagi, T. (2012). Fungal denitrification and nitric oxide reductase cytochrome P450nor. Philosophical Transactions of the Royal Society B: Biological Sciences, 367(1593), 1186–1194. https://doi.org/10.1098/rstb.2011.0335 Sithtisarn, S., Yokthongwattana, K., Mahong, B., Roytrakul, S., Paemanee, A., Phaonakrop, N., & Yokthongwattana, C. (2017). Comparative proteomic analysis of Chlamydomonas reinhardtii control and a salinity-tolerant strain revealed a differential protein expression pattern. Planta, 246(5), 843–856. https://doi.org/10.1007/s00425-017-2734-4 Sivakumar, P., Sharmila, P., & Pardha Saradhi, P. (2000). Proline alleviates salt-stress-induced enhancement in ribulose-1,5-bisphosphate oxygenase activity. Biochemical and Biophysical Research Communications, 279(2), 512–515. https://doi.org/10.1006/bbrc.2000.4005 Steinbeck, J., Ross, I. L., Rothnagel, R., Gäbelein, P., Schulze, S., Giles, N., … Hankamer, B. (2018). Structure of a PSI-LHCI-cyt b6f supercomplex in Chlamydomonas reinhardtii promoting cyclic electron flow under anaerobic conditions. Proceedings of the National Academy of Sciences of the United States of America, 115(41), 10517–10522. https://doi.org/10.1073/pnas.1809973115 Strand, D. D., Livingston, A. K., Satoh-Cruz, M., Froehlich, J. E., Maurino, V. G., & Kramer, D. M. (2015). Activation of cyclic electron flow by hydrogen peroxide in vivo. Proceedings of

209

the National Academy of Sciences of the United States of America, 112(17), 5539–5544. https://doi.org/10.1073/pnas.1418223112 Subramanyam, R., Jolley, C., Thangaraj, B., Nellaepalli, S., Webber, A. N., & Fromme, P. (2010). Structural and functional changes of PSI-LHCI supercomplexes of Chlamydomonas reinhardtii cells grown under high salt conditions. Planta, 231(4), 913–922. https://doi.org/10.1007/s00425-009-1097-x Sudhir, P., & Murthy, S. D. S. (2004). Effects of salt stress on basic processes of photosynthesis. Photosynthetica, 42(4), 481–486. https://doi.org/10.1007/S11099-005-0001-6 Suorsa, M. (2015). Cyclic electron flow provides acclimatory plasticity for the photosynthetic machinery under various environmental conditions and developmental stages. Frontiers in Plant Science, 6(September), 1–8. https://doi.org/10.3389/fpls.2015.00800 Szyszka-Mroz, B., Cvetkovska, M., Ivanov, A. G., Smith, D. R., Possmayer, M., Maxwell, D. P., & Hüner, N. P. A. (2019). Cold-adapted protein kinases and thylakoid remodeling impact energy distribution in an antarctic psychrophile. Plant Physiology, 180(3), 1291–1309. https://doi.org/10.1104/pp.19.00411 Szyszka-Mroz, B., Pittock, P., Ivanov, A. G., Lajoie, G., & Hüner, N. P. A. (2015). The antarctic psychrophile Chlamydomonas sp. UWO 241 preferentially phosphorylates a photosystem I- cytochrome b6/f supercomplex. Plant Physiology, 169(1), 717–736. https://doi.org/10.1104/pp.15.00625 Szyszka, B., Ivanov, A. G., & Hüner, N. P. A. (2007). Psychrophily is associated with differential energy partitioning, photosystem stoichiometry and polypeptide phosphorylation in Chlamydomonas raudensis. Biochimica et Biophysica Acta - Bioenergetics, 1767(6), 789– 800. https://doi.org/10.1016/j.bbabio.2006.12.001 Takahashi, H., Clowez, S., Wollman, F. A., Vallon, O., & Rappaport, F. (2013). Cyclic electron flow is redox-controlled but independent of state transition. Nature Communications, 4. https://doi.org/10.1038/ncomms2954 Takahashi, H., Iwai, M., Takahashi, Y., & Minagawa, J. (2006). Identification of the mobile light- harvesting complex II polypeptides for state transitions in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America, 103(2), 477–482. https://doi.org/10.1073/pnas.0509952103

210

Takahashi, H., Schmollinger, S., Lee, J. H., Schroda, M., Rappaport, F., Wollman, F. A., & Vallon, O. (2016). PETO Interacts with Other Effectors of Cyclic Electron Flow in Chlamydomonas. Molecular Plant, 9(4), 558–568. https://doi.org/10.1016/j.molp.2015.12.017 Terashima, M., Petroutsos, D., Hüdig, M., Tolstygina, I., Trompelt, K., Gäbelein, P., … Hippler, M. (2012). Calcium-dependent regulation of cyclic photosynthetic electron transfer by a CAS, ANR1, and PGRL1 complex. Proceedings of the National Academy of Sciences of the United States of America, 109(43), 17717–17722. https://doi.org/10.1073/pnas.1207118109 Valledor, Luis; Weckwerth, W. (2014). Chapter 25. In Plant Proteomics: Methods and Protocols, Methods in Molecular Biology, (Vol. 1072, pp. 347–358). https://doi.org/10.1016/b978-0-08- 011049-3.50031-5 Van Lis, R., Popek, M., Couté, Y., Kosta, A., Drapier, D., Nitschke, W., & Atteia, A. (2017). Concerted up-regulation of aldehyde/alcohol dehydrogenase (ADHE) and starch in Chlamydomonas reinhardtii increases survival under dark anoxia. Journal of Biological Chemistry, 292(6), 2395–2410. https://doi.org/10.1074/jbc.M116.766048 Voss, I., Sunil, B., Scheibe, R., & Raghavendra, A. S. (2013). Emerging concept for the role of photorespiration as an important part of abiotic stress response. Plant Biology, 15(4), 713– 722. https://doi.org/10.1111/j.1438-8677.2012.00710.x Wang, N., Qian, Z., Luo, M., Fan, S., Zhang, X., & Zhang, L. (2018). Identification of salt stress responding genes using transcriptome analysis in green alga Chlamydomonas reinhardtii. International Journal of Molecular Sciences, 19(11). https://doi.org/10.3390/ijms19113359 Wang, X., Liu, W., Xin, C., Zheng, Y., Cheng, Y., Sun, S., … Yuan, J. S. (2016). Enhanced limonene production in cyanobacteria reveals photosynthesis limitations. Proceedings of the National Academy of Sciences of the United States of America, 113(50), 14225–14230. https://doi.org/10.1073/pnas.1613340113 Wang, Y., Stessman, D. J., & Spalding, M. H. (2015). The CO2 concentrating mechanism and photosynthetic carbon assimilation in limiting CO2: How Chlamydomonas works against the gradient. Plant Journal, 82(3), 429–448. https://doi.org/10.1111/tpj.12829 Wingler, A., Lea, P. J., Quick, W. P., & Leegood, R. C. (2000). Photorespiration: Metabolic pathways and their role in stress protection. Philosophical Transactions of the Royal Society B: Biological Sciences, 355(1402), 1517–1529. https://doi.org/10.1098/rstb.2000.0712

211

Yamori, W., & Shikanai, T. (2016). Physiological Functions of Cyclic Electron Transport Around Photosystem i in Sustaining Photosynthesis and Plant Growth. Annual Review of Plant Biology, 67(July), 81–106. https://doi.org/10.1146/annurev-arplant-043015-112002 Yang, X., Li, Y., Chen, H., Huang, J., Zhang, Y., Qi, M., … Li, T. (2020). Photosynthetic response mechanism of soil salinity-induced cross-tolerance to subsequent drought stress in tomato plants. Plants, 9(3), 1–15. https://doi.org/10.3390/plants9030363 Yi, S. Y., Ku, S. S., Sim, H. J., Kim, S. K., Park, J. H., Lyu, J. Il, … Liu, J. R. (2017). An alcohol dehydrogenase gene from synechocystis sp. Confers salt tolerance in transgenic tobacco. Frontiers in Plant Science, 8(November), 1–13. https://doi.org/10.3389/fpls.2017.01965 Ziotti, A. B. S., Silva, B. P., Sershen, & Lima Neto, M. C. (2019). Photorespiration is crucial for salinity acclimation in castor bean. Environmental and Experimental Botany, 167(May), 103845. https://doi.org/10.1016/j.envexpbot.2019.103845

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4.6 APPENDIX

A UWO241 B ICE-MDV C C.reinhardtii

0.8 0.8 0.8

0.6 0.6 0.6

0.4 0.4 MAX 0.4 MAX MAX F F F

0.2 0.2 0.2

0.0 0.0 0.0

LS-StateLS-State I IIHS-StateHS-State I II LS-STATE I HS-STATE I LS-STATE I LS-STATE II HS-STATE II LS-STATEHS-STATE II HS-STATE I II

Appendix Figure 4.1: PSII state transition test for the three Chlamydomonas spp under low (LS) and high (HS) salinity. The maximum PSII fluorescence values (FMAX) are shown for all three strains (UWO241- A, ICE-MDV-B and C. reinhardtii-C) under state I and state 2 conditions. State I: DCMU, State II: FCCP. (n=4-6, dotted line=mean value)

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A B C 1.0 UWO241 LS 1.0 1.0 ICE-MDV LS C. reinhardtii LS UWO241 HS ICE-MDV HS 0.8 0.8 0.8 C. reinhardtii HS 0.6 0.6 0.6

0.4 0.4 0.4

Absorbance Absorbance Absorbance 0.2 0.2 0.2

0.0 0.0 0.0 0 2000 4000 6000 0 2000 4000 6000 0 2000 4000 6000 time(ms) time (ms) time (ms)

Appendix Figure 4.2: P700 reduction kinetics of the three Chlamydomonas species under low and high salinity. A. UWO241. B. ICE-MDV. C. C. reinhardtii. Low salinity (LS) traces are shown in blue. High salinity (HS) traces are shown in red.

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CHAPTER 5

CONCLUSION

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CHAPTER V. CONCLUSION

Climate change is expected to negatively impact crop productivity in the future, and it’s predicted that the major food producing crops will decrease their productivity by 50%. In addition, the food demands of the population are expected increase several folds by 2050 (Alexandratos & Brunismas, 2012). Thus, there is a need for understanding how photosynthetic organisms cope with environmental change and which processes are predominant under those conditions. Green algae possess similar photosynthetic machinery as land plants and are excellent model to study the impact of environmental stresses on the photosynthesis. One of the major abiotic stress that photosynthetic organisms deal with is salinity stress (Negrão et al., 2017). Soil salinity results in damage to many food producing crops yearly and this phenomenon is expected to exacerbate with climate change. Photosynthesis consists of two major electron transport pathways: Linear electron flow (LEF) and cyclic electron flow (CEF) (Yamori & Shikanai, 2016). The latter has been shown to be essential for balancing ATP:NADPH ratio for carbon fixation and is especially upregulated under conditions of stress (Kramer & Evans, 2011). During stress, CEF helps in production of excess ATP for carbon fixation and downstream metabolism, and/or quenching of excess energy as non-photochemical quenching (Yamori & Shikanai, 2016). CEF is a ubiquitous mechanism operating in all photosynthetic organism and thus forms an excellent target to improve photosynthetic performance during stress. Although the role of CEF has been studied in the context of short-term transient stress (H. Chen et al., 2015; Takahashi et al., 2016; Terashima et al., 2012), not much is known on the role of CEF during long-term stress acclimation. This is especially important in the context of climate change as the organisms are predicted to deal with long-term steady state stress in their environment. Under short-term stress, CEF has been shown to be associated with formation of PSI-supercomplexes, that help in increasing the efficiency of electron transfer around PSI (Iwai et al., 2010; Terashima et al., 2012). However, it is not understood whether such supercomplexes operate under steady-state long-term stress conditions. In this study, we have used three different Chlamydomonas species, with a range salinity tolerance to understand the role of CEF and PSI-supercomplexes in long-term salinity acclimation. In addition, we attempted to dissect the effect of increased CEF associated salinity

216 acclimation on downstream metabolism of a psychrophilic and mesophilic salt-tolerant green algae. Chlamydomonas sp. UWO241 is a halotolerant psychrophilic green alga isolated from ice-covered Antarctic lake (Dolhi et al., 2013). Past studies have described ways in which UWO241 has modified its photosynthetic apparatus to deal with the permanent stress of low temperature and hypersalinity (R. Morgan-Kiss et al., 2002; Morgan et al., 1998; Szyszka-Mroz et al., 2015; Szyszka et al., 2007). This project focused on its ability to maintain high rates of CEF in high salinity conditions. There was a gap in the knowledge of the downstream metabolism that is supported by the organism’s altered physiology of increased CEF. In this study, we attempted to understand the outcomes of increased CEF in UWO241 and analyzed the whole cell proteome of UWO241 under low and high salinity conditions to identify the metabolic pathways that maybe differentially upregulated in those conditions. We found that UWO241 has an altered photosynthetic apparatus that helps in constitutive upregulation of CEF under both low and high salinity conditions, and that a stable PSI-supercomplex is associated with CEF under hypersalinity. This increased CEF under high salinity conditions was shown to help in excess ATP production that feeds into downstream carbon fixation and secondary metabolism. In addition, we observed upregulation of glycerol and starch biosynthesis. We suggest the UWO241 possesses an altered photosynthetic apparatus that helps rewire the downstream metabolism to cope with low temperature and hypersalinity. To further dissect the role of CEF in long-term stress acclimation, we moved to the genetically described model C. reinhardtii. Our goal was to evolve the strain under autotrophic conditions to understand its photosynthetic response to long-term salinity stress as well as develop a strain with high rates of CEF and a stable supercomplex for future studies. Previous studies had shown evolution of C. reinhardtii under salinity stress led to downregulation of photosynthesis and preferential utilization of acetate (Perrineau et al., 2014). We predicted that evolution of C. reinhardtii without external carbon source would force the organism to rely on photosynthesis for salinity acclimation and CEF may be essential for this acclimation. We found that the evolved strain was fast growing and had sustained-high rates of CEF like UWO241 along with constitutive capacity for non-photochemical quenching. In addition, the evolved strain also maintained high rates of oxygen evolution and respiration. Our proteome analysis showed that the evolved strain has downregulated energy expensive processes such as

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photorespiration but has upregulated ATP-generating mitochondrial respiration. In addition, we found that evolved strain upregulates transient starch metabolism, which may feed into glycerol and ethanol biosynthesis, providing stored carbon reserves for robust growth. We thus suggest that CEF is important for long-term salinity acclimation and may help in rewiring the downstream metabolism towards upregulation of osmolytes and storage carbon. Further studies are needed to dissect the outcome of constitutive upregulation of CEF during salinity stress acclimation. PSI-supercomplexes have been widely studied under short-term stresses such as anoxia and high light stress (state 2 conditions) (Z. Huang et al., 2021; Steinbeck et al., 2018), however its formation and composition during long-term stress acclimation is not understood. In addition, short-term stress response such as state transition are downregulated in UWO241 that displays increased CEF and forms PSI-supercomplex under high salinity. In this study, we attempted to understand whether increased CEF and PSI-supercomplex formation are associated with long- term salinity acclimation in three different Chlamydomonas species: salt-resistant UWO241, moderately salt-tolerant Antarctic C. sp. ICE-MDV and salt-sensitive C. reinhardtii. We also measured the propensity for state transition after salinity acclimation in all the three species We observed that all three species had increased rates of CEF after salinity acclimation, and that the two Antarctic species displayed fastest CEF rates, possibly as a result of additional low- temperature acclimation. Moreover, state transition response was attenuated after long-term salinity acclimation. We also show association of PSI-supercomplexes with high CEF displaying salinity acclimated cultured from all three species. Lastly, the high salt supercomplex of C. reinhardtii shared many proteins with the widely studied state 2 supercomplex (Fig. 5.1). The HS-supercomplex was shown to possess many proteins involved in calcium homeostasis, as well as reduced abundance of LHCI proteins. In conclusion, we show that CEF and PSI-supercomplexes are not only important for short-term stress, but also during long-term salinity stress acclimation. The high salt supercomplex shares many similarities with state 2 supercomplex and appear to be species specific rather than stress specific. In addition, increased rates of CEF were shown to be associated with rewired downstream metabolism in both halotolerant UWO241 and salinity- evolved C. reinhardtii in high salinity (Fig. 5.1). This rewired metabolism helped in upregulation

218 of carbon fixation, secondary metabolism and glycerol biosynthesis in UWO241. On the other hand, transient starch metabolism, ethanol and glycerol biosynthesis processes were upregulated

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in Acclimationthe evolved strain. Adaptation Salinity stress

Wildtype C. reinhardtii Evolved C. reinhardtii UWO241

• Constitutive CEF • Constitutive CEF • Cyclic electron flow

• Thermal dissipation not induced • Constitutive thermal dissipation • Constitutive thermal dissipation

• Supercomplex • Supercomplex • Possible increased ATP from: formation formation ATPase Fd FNR • Increased CEF FtsH Fd • Increased Respiration FtsH Cytbf PSI • Decreased Photorespiration Cytbf PSI Lhca LHCII

CAS Lhca3 PETO CAS PC PsbP PC PsbP • Loss of state transitions • Restructured downstream • Short-term state transition response metabolism for robust growth Metabolism Excess ATP Carbon fixation Transient starch biosynthesis Starch metabolism Storage compound biosynthesis ATP Transport proteins secondary metabolism ATP Oxidative stress proteins ATP

Figure 5.1. Photosynthetic and metabolic changes associated with acclimation and adaptation to salinity stress in green algae Chlamydomonas reinhardtii and UWO241. The arrow represents amount of time the alga is exposed to salinity stress. Results from studies on wiltype and evolved C. reinhardtii represent acclimation to 6 days and 24 months to salinity stress respectively. UWO241 is a halophyte that is adapted to and survives in its natural environment with high concentrations of NaCl. Predicted models of supercomplex are shown. Thermal dissipation: Non-photochemical quenching (NPQ)

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REFERENCES

Alexandratos, N., & Brunismas, J. (2012). WORLD AGRICULTURE TOWARDS 2030/2050. JESA Working Papers 12-03. https://doi.org/10.22004/ag.econ.288998 Chen, H., Hu, J., Qiao, Y., Chen, W., Rong, J., Zhang, Y., … Wang, Q. (2015). Ca 2+ -regulated cyclic electron flow supplies ATP for nitrogen starvation-induced lipid biosynthesis in green alga. Scientific Reports, 5(October), 1–12. https://doi.org/10.1038/srep15117 Dolhi, J. M., Maxwell, D. P., & Morgan-Kiss, R. M. (2013). Review: The Antarctic Chlamydomonas raudensis: An emerging model for cold adaptation of photosynthesis. Extremophiles, 17(5), 711–722. https://doi.org/10.1007/s00792-013-0571-3 Huang, Z., Liangliang, S., Wenda, W., Zhiyuan, M., Xiaohan, Y., Tingyun, K., … Guangye, H. (2021). Structure of photosystem I-LHCI-LHCII from the green alga Chlamydomonas reinhardtii in State 2. Nature Communications, (12(1)), 1–14. https://doi.org/10.1038/s41467- 021-21362-6 Iwai, M., Takizawa, K., Tokutsu, R., Okamuro, A., Takahashi, Y., & Minagawa, J. (2010). Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis. Nature, 464(7292), 1210–1213. https://doi.org/10.1038/nature08885 Kramer, D. M., & Evans, J. R. (2011). The importance of energy balance in improving photosynthetic productivity. Plant Physiology, 155(1), 70–78. https://doi.org/10.1104/pp.110.166652 Morgan-Kiss, R., Ivanov, A. G., Williams, J., Khan, M., & Huner, N. P. A. (2002). Differential thermal effects on the energy distribution between photosystem II and photosystem I in thylakoid membranes of a psychrophilic and a mesophilic alga. Biochimica et Biophysica Acta - Biomembranes, 1561(2), 251–265. https://doi.org/10.1016/S0005-2736(02)00352-8 Morgan, R. M., Ivanov, A. G., Priscu, J. C., Maxwell, D. P., & Huner, N. P. A. (1998). Structure and composition of the photochemical apparatus of the Antarctic green alga, Chlamydomonas subcaudata. Photosynthesis Research, 56(3), 303–314. https://doi.org/10.1023/A:1006048519302 Negrão, S., Schmöckel, S. M., & Tester, M. (2017). Evaluating physiological responses of plants to salinity stress. Annals of Botany, 119(1), 1–11. https://doi.org/10.1093/aob/mcw191 Perrineau, M. M., Zelzion, E., Gross, J., Price, D. C., Boyd, J., & Bhattacharya, D. (2014).

221

Evolution of salt tolerance in a laboratory reared population of Chlamydomonas reinhardtii. Environmental Microbiology, 16(6), 1755–1766. https://doi.org/10.1111/1462-2920.12372 Steinbeck, J., Ross, I. L., Rothnagel, R., Gäbelein, P., Schulze, S., Giles, N., … Hankamer, B. (2018). Structure of a PSI-LHCI-cyt b6f supercomplex in Chlamydomonas reinhardtii promoting cyclic electron flow under anaerobic conditions. Proceedings of the National Academy of Sciences of the United States of America, 115(41), 10517–10522. https://doi.org/10.1073/pnas.1809973115 Szyszka-Mroz, B., Pittock, P., Ivanov, A. G., Lajoie, G., & Hüner, N. P. A. (2015). The antarctic psychrophile Chlamydomonas sp. UWO 241 preferentially phosphorylates a photosystem I- cytochrome b6/f supercomplex. Plant Physiology, 169(1), 717–736. https://doi.org/10.1104/pp.15.00625 Szyszka, B., Ivanov, A. G., & Hüner, N. P. A. (2007). Psychrophily is associated with differential energy partitioning, photosystem stoichiometry and polypeptide phosphorylation in Chlamydomonas raudensis. Biochimica et Biophysica Acta - Bioenergetics, 1767(6), 789– 800. https://doi.org/10.1016/j.bbabio.2006.12.001 Takahashi, H., Schmollinger, S., Lee, J. H., Schroda, M., Rappaport, F., Wollman, F. A., & Vallon, O. (2016). PETO Interacts with Other Effectors of Cyclic Electron Flow in Chlamydomonas. Molecular Plant, 9(4), 558–568. https://doi.org/10.1016/j.molp.2015.12.017 Terashima, M., Petroutsos, D., Hüdig, M., Tolstygina, I., Trompelt, K., Gäbelein, P., … Hippler, M. (2012). Calcium-dependent regulation of cyclic photosynthetic electron transfer by a CAS, ANR1, and PGRL1 complex. Proceedings of the National Academy of Sciences of the United States of America, 109(43), 17717–17722. https://doi.org/10.1073/pnas.1207118109 Yamori, W., & Shikanai, T. (2016). Physiological Functions of Cyclic Electron Transport Around Photosystem i in Sustaining Photosynthesis and Plant Growth. Annual Review of Plant Biology, 67(July), 81–106. https://doi.org/10.1146/annurev-arplant-043015-112002

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